Recombinant protein–based medicinal products and modern cell-based vaccines have a very strong safety history with respect to viral and microbial contamination. However, virus contamination incidents do occur occasionally in manufacturing processes, and they can consume many resources and be expensive to rectify.

The root cause of contamination incidents in recent years is most likely the use of contaminated raw materials. These include bovine serum contaminated with reovirus, epizootic hemorrhagic disease virus, Cache valley virus or vesivirus 2117; porcine trypsin contaminated with porcine circovirus; and other media components contaminated with minute virus of mice (MVM).

In those cases, no virus was detected through routine raw-materials screening because of limitations either in the sensitivity of assays used or in the amount of material screened. Components of some materials (such as antivirus antibodies that may be present in bovine serum) can also inhibit virus detection. Metagenomic techniques (such as massively parallel sequencing or virus nucleic acid chips) have detected nucleic acid sequences for new viruses that may be more prevalent in serum batches than such viruses that are detected in classical quality control assays used by serum suppliers (1, 2).

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The European Medicines Agency (EMA) issued two guidance documents in 2013 addressing the production and quality control of bovine serum and porcine trypsin. At the end of May, the agency issued a revision to its guideline on the use of bovine serum in manufacturing human biological medicinal products (3). And in March, the EMA issued a new draft guideline on the use of porcine trypsin used in the manufacture of human biological medicinal products (4).

The revised bovine serum guidance, effective on 1 December 2013, covers the types and sources of serum, preparation of batches, certificate of analysis, testing for adventitious agents, tests for toxicity-cell growth, viral inactivation, and dossier requirements for marketing authorization/variations. Specified viruses for which infectivity assays are required in the EMA guidance are similar to those required in the 9 CFR test. If an infectious virus is detected in a batch of serum, then that batch should not be used in the manufacture of a medicinal product, with the exception of bovine viral diarrhea virus (BVDV). Serum should be tested before any viral inactivation treatment is performed.

Because BVDV is a highly prevalent infection in cattle, and the presence in serum cannot be completely eliminated from all processes, it is possible to use this serum as long as the titer of BVDV is below the level that has been shown to be effectively inactivated. To ensure that BVDV can be detected, serum should be screened for the presence of anti-BVDV antibodies. The first major change in the revised guideline is removal of the allowable limit of no more than two logs of BVDV infectivity. Manufacturers should assess the effects of anti-BVDV antibodies on their ability to reveal any residual virus in the product after inactivation.

The guidance document does not mention testing for the presence of neutralizing antibodies to any other virus. Antibodies to vesivirus 2117 (which has caused contamination events at both European and US manufacturing sites) have been detected in the serum from asignificant percentage of USA cattle (5). That could possibly mask the detection of the virus.

The original guidance document suggested that manufacturers should also test serum for the presence of infectious bovine polyoma virus (BPyV), which has been detected by polymerase chain reaction (PCR) in most batches of bovine serum. Following comments to the guideline from serum suppliers, the EMA added a footnote indicating that, for the time being, such testing is not required. The agency claims that infectivity assays are difficult to interpret and not widely available. The agency indicates that this guidance will be reviewed if further data become available or a contamination event occurs.

Nonetheless, the guidance document still indicates that the EMA expects manufacturers to be aware of emerging viruses and take appropriate action to detect them. In recent years, genetic material from a number of new viruses has been detected in bovine serum by massively parallel sequencing: notably bovine parvovirus 2 and 3, bovine adeno associated virus 2, bovine norovirus, and bovine kobuvirus. The genomes for those viruses are present in a high proportion of batches, but infectivity assays for them have yet to be developed.

Following detection of porcine circovirus contamination in two rotavirus vaccines (6) — probably originating from contaminated porcine trypsin — the EMA announced in 2012 that it would draft a guidance document on the use of porcine trypsin. That draft guidance was published on 1 March for consultation until 31 August 2013 (4). It covers testing for adventitious agents, manufacture, validation of virus-reducing capacity of a manufacturing process, quality controls, use of alternative reagents, and risk assessment.

Although trypsin should be sourced only from pigs that are fit for human consumption, contamination of animal-derived material with infectious virus still poses a risk. So pooled starting material should be tested before any virus inactivation/ removal steps. Testing of individual pancreatic glands, however, is not considered possible for economic and organizational reasons. Because frozen pancreatic glands are usually directly extracted into alcohol-containing solutions, the guidance does not define the exact point of the process at which to test for infectious virus. That point should be determined and justified.

A large number of porcine viruses have the potential to infect humans or replicate in human and primate cells. So a general in vitro virus assay should be performed using two detector cell lines (Vero and porcine cells). Virus replication would be detected through viral cytopathic effect and/or hemadsorbtion. The guidance suggests that specific tests for porcine viruses that are not detected by a general cell culture assay should also be considered following a product-specific risk analysis. PCR assays for stable viruses such as porcine circovirus and hepatitis E are examples of specific tests that should be considered for all trypsin batches.

Trypsin manufacturers should incorporate two complementary virus-reduction steps into their manufacturing processes unless otherwise justified. Examples of effective virus-inactivating steps are low pH treatment and gamma or UV-C irradiation. Such steps should be validated as described in the EMA guidance on virus validation studies (7). In addition, trypsin manufacturers should incorporate validated cleaning measures to minimize the risk of batch-to-batch cross contamination with infectious viruses.

The guideline recommends use of recombinant or plant-derived trypsin but acknowledges that such alternatives must be assessed for their performance characteristics. If a manufacturer intends to use porcine trypsin in the
manufacture of medicinal products, it is expected that the company would analyze associated risks using the principles outlined in European Pharmacopoeia 5.1.7 Viral Safety. This assessment would cover the sourcing and manufacture of the trypsin and its use in manufacturing processes. The guideline is for prospective implementation. However, in the light of reported contamination events, the EMA recommends that manufacturers reassess the virus safety of authorized live-virus vaccines that use porcine trypsin in their manufacturing processes.

These two EMA guidelines define expected approaches that should be taken to minimize the risk of virus contamination through use of the animal-derived raw materials described above. Industry and regulators are also concerned about contamination events — such as MVM contamination of Chinese hamster ovary (CHO) cells — that can arise through contamination of other raw materials or components of cell culture media. Companies are evaluating the potential of incorporating virus-inactivating steps into their manufacturing processes. Strategies include gamma irradiation, UV-C irradiation, high-temperature– short-time (HTST) treatment of raw materials, and the use of virus-removal filters to reduce the risk of virus contamination in raw materials. Although the use of those processes are not described in any recent regulatory guideline, their implementation is in line with the expectations detailed in ICH Q5Athat potential viral contamination should be controlled by the selection and testing of raw materials, including media components, for the absence of viral contaminants.

Author Details
Dr. Martin Wisher is senior director of regulatory affairs at BioReliance Ltd., Todd Campus, West of Scotland Science Park, Glasgow, G20 0XA, Scotland; 44-141-946- 9999, fax 44-141-946-0000; martin.wisher@bioreliance.com; www.bioreliance.com.

REFERENCES

The post Virus Risk Mitigation for Raw Materials appeared first on BioProcess International.

Food and Drug Administration (FDA) and European good manufacturing practices (GMPs) require integrity testing of sterilizing-grade filters for producing injectables and other biologics. The diffusion test (also called the forward-flow test) and bubble-point test (also called the disk test) of a sterilizing-grade filter are both filter-integrity tests. The accuracy of both relies on calibration of a pressure sensor in the respective integrity test unit.

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Calibration of the pressure sensor of a filter-integrity testing device is an essential part of quality assurance in lot release. If yearly calibration reveals a pressure reading offset outside specifications, then a common conclusion is that the accuracy of all integrity test results from the past year must be questioned. Generally, for a maximum allowable offset of 9 mbar, for example, all test results since the previous calibration are considered reliable if the yearly calibration shows an offset of ≤9 mbar. If the yearly calibration shows an offset >9 mbar — even by only 1 mbar — then all test results since the previous calibration are considered questionable.

PRODUCT FOCUS: PARENTERAL PRODUCTS
PROCESS FOCUS: PURIFICATION, STERILE FILTRATION, QUALITY ASSURANCE
WHO SHOULD READ: PROCESS DEVELOPMENT AND MANUFACTURING
KEYWORDS: INJECTABLES, STERILIZING-GRADE FILTRATION, INTEGRITY TESTING, CALIBRATION
LEVEL: ADVANCED

The limitation of that approach is that the real impact is not evaluated. In some cases, the influence of a calibration offset is negligible. In other cases, the calibration offset overestimates the diffusion (forward flow) value, thereby generating no risk for false conformity test results. Quality assurance divisions need an appropriate factual tool to estimate the actual impact of a calibration offset on integrity test results.

Here I describe an analytical approach of an event-based pressure reading offset based on theory and experimental trials. The method can prevent unnecessary time-consuming quality assurance investigations. It must be adapted to the type of filter-integrity test unit used because of different technological approaches and algorithms.

A diffusion test measures the amount of gas that — when a given gas pressure is applied on the upstream side of a wetted membrane filter cartridge — dissolves into the wetting liquid and diffuses through the wetted membrane. Capillary forces keep the wetting liquid from being expelled from membrane pores. A filter is considered being integer when the measured diffusion value does not exceed the validated maximum value.

A multipoint diffusion test measures diffusion values at predefined pressures. A diffusion profile is a function of applied gas pressure on the upstream side of a wetted membrane filter cartridge. A multipoint diffusion test is not a common integrity test. Rather, it is typically used for validation to evaluate characteristics of a diffusion profile.

A bubble-point test measures a gradually increasing diffusion rate of a filter cartridge at stepwise higher differential gas pressures. The bubble point is detected when gas flow goes from a diffusion value at a given pressure step p_n (pores still filled with wetting liquid) to a bulk flow at p_n+1 (biggest pores no longer filled with water). That is characterized by an exponentially increasing gas flow rate through the filter cartridge. An algorithm is used to detect this phenomenon. A filter is considered being integer when the measured bubble point equals or exceeds the validated minimum bubble point.

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Figure 1:  ()

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Figure 2:  ()

Multipoint diffusion tests were performed on a filter capsule (diffusion tests at increasing pressure levels within the same test sequence). Tests involved the same filter (Figure 3 test setup) using a calibrated integrity test unit. Programmed test pressures were 2,100, 2,300, 2,500, and 2,700 mbar to verify the effect on the test value of the actual test pressure on a given filter. No reference instruments were used. The test was repeated twice for a total of three multipoint diffusion tests.

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Figure 3:  ()

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Table 1:&#8 194; ()

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Table 2:&#8 194; ()

As a second step, diffusion test accuracies of two integrity test units were verified compared with a reference method. The first series of diffusion accuracy measurements (Figure 4 test setup) were conducted when the integrity test units were correctly calibrated. The second series of diffusion accuracy measurements were made after having generated different degrees of pressure reading offsets: 50 mbar full scale on one integrity-test unit; 100 mbar full scale on one integrity-test unit; 200 mbar full scale on both integrity-test units. Such pressure reading offsets are extremely high and may not occur spontaneously in reality. Nevertheless, those extreme settings were necessary for obtaining quantifiable and interpretable test data.

Researchers decided that having an nonparallel pressure reading offset with different atmospheric reading (Figure 2D) was the most representative offset and that more information could be achieved than when having a parallel offset (Figure 2A). The worst case was when the test unit overestimates the test pressure, thereby applying a test pressure that is too low.

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Figure 4:  ()

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Figure 5:  ()

As the third step, bubble-point tests were performed on a filter capsule using a calibrated integrity-test unit (Figure 3 setup). One additional series of bubble-point tests was performed on the same filter capsule using the same integrity-test unit, but with a pressure reading offset of 200 mbar full scale. Reference instruments were not used. These tests were repeated for a total of three tests with the calibrated unit and three tests with a 200-mbar pressure reading offset full scale.

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Table 3:&#8 194; ()

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Table 4:&#8 194; ()

We used the following materials: integrity-test units (Sartocheck 4 v2.03 serial number 17101314 and 17201162); reference manometer GE Drück DPI 150 (absolute and relative pressure reading); and reference gas flow meter Bronkhorst F111 CAAD-22-V. We used a reference temperature sensor Sika MH 3710 and GTF-401 brands. Stainless steel volume and tubings were from Sartorius Stedim, and the filter capsule was a Sartopore 2 Maxicap 0.2 µm (lot 482503, individual number 128).

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Table 5:&#8 194; ()

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Table 6:&#8 194; ()

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Table 7:&#8 194; ()

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Figure 6:  ()

Tables 234567 show the results obtained for all tests. Table 2 lists data from a multipoint diffusion test of a filter capsule using a calibrated integrity test unit. Tables 34567 show results from calibrated and decalibrated tests for the Sartopore 4 integrity-test unit compared with a flow reference.

The influence on the diffusion test value from a pressure reading offset comes from two factors. First, the difference between the displayed test pressure and the actual applied test pressure reduces or increases the driving force of the diffusion test. The results obtained in test 1 (multipoint diffusion test in Table 2) show the effect of test pressure on diffusion test value. Second, the difference in pressure-reading slope affects the pressure drop reading. The results obtained in test 2 (compared with those from a flow reference meter in Tables 34567) show the effect of pressure-reading slope (pressure drop measurement) on diffusion test value. Both factors then must be combined to estimate the cumulative influence on the diffusion test value.

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Figure 7:  ()

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Table 8:&#8 194; ()

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Table 9:&#8 194; ()

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Table 10:&#8 194; ()

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Table 11:&#8 194; ()

Effect of Deviation in Applied Test Pressure: The influence on the diffusion test value is directly proportional to the deviation of the applied test pressure according to Fick’s law (
Equation 1). In that equation, N is the diffusive flux of the test gas, D is the diffusivity of the test gas through the wetting liquid, H is the solubility coefficient of the test gas in the wetting liquid, p is the applied differential pressure, ϕ is the overall porosity of the structure, and L is thickness of the wet layer. Fick’s law is explained in PDA Technical Report 26 (1).

All parameters for a given filter are fixed, so the volumetric flow can be reduced to
Equation 2. F is the volumetric diffusive flow, and K₁ is the proportionality constant (slope). This equation should be true within a certain range from traditional diffusion test pressure depending on a filter cartridge’s membrane configuration. As Figure 5 shows, linearity ceases to exist when the overproportional bubble-point region has been reached (when the wetting liquid is pressed out from the biggest pores). According to Fick’s law, the effect (as a percentage) on the diffusion value (ΔD) due to a deviation in the applied test pressure (p_Δ) during a diffusion test can then be expressed using
Equation 3, in which p_test is the normally applied test pressure (mbar relative). Taking an actual pressure reading offset of 200 mbar at 2,500 mbar of differential test pressure, the theoretical value of the pressure deviation influence on the diffusion value is as shown in
Equation 4.

The experimental value based on an average of three multipoint diffusion test measurements on the same capsule was +7.9% for +200 mbar and –7.9% for –200 mbar (Table 2). To get an actual pressure reading offset of 200 mbar at a test pressure of 2,500 mbar — if using the same type of nonparallel offset as in Ta
ble 1 — the full scale (9,500 mbar absolute) offset would have to have been 543 mbar.

Theoretical and experimental results match within the 200-mbar range. The three tests with a 400-mbar test pressure offset (2,100 mbar instead of 2,500 mbar showed a deviation of –13% instead of the expected –16%). So we used this theoretical calculation for pressure reading offsets, which are ≤8% of the programmed test pressure.

Equation 1:

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Equation 2:

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Equation 3:

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Equation 4:

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Equation 5:

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Equation 6:

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Equation 7:

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Equation 8:

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Equation 9:

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Equation 10:

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Equation 11:

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Equation 12:

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Equation 13:

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Equation 14:

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Equation 15:

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Equation 16:

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Equation 17:

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Equation 18:

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Figure 8:  ()

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Figure 9:  ()

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Table 12:&#8 194; ()

Calculation of Actual Initial Starting Pressure based on Calibration Values: A perfect calibration curve (g_α) can be represented as a straight line with slope = 1 and slope intercept = 0, leading to the equation g_α = 1x, where x is the real pressure and “y” the pressure read by the integrity test unit. A pressure reading offset curve (g_β) as generated for these trials can also be represented as a straight line but with a different slope (m) and the slope intercept (b) as in
Equation 5.

The deviation in pressure reading at a certain reference pressure p₁ (here called x₁) is illustrated by ▵y₁ (Figure 6) and expressed by
Equation 6.

In the case of a calibration offset, the interesting portion of the calibration curve are the calibration points p_A (the calibration point above p₁ — the initial test pressure) and p_B (the calibration point below p₂ — the pressure at the end of the test) (Figure 7). That means that even if the rest of the pressure deviation values do not form a straight line, the section p_A to p_B is enough for calculating the impact of the pressure reading deviation on the test pressure.

Equation 7 is used to calculate slope m of the pressure reading curve at the appropriate pressure interval. The slope intercept “b” can be calculated using
Equation 8. Based on ▵y = (m – 1)x₁ + b, the pressure reading offset at the programmed test pressure, p₁ can then be calculated by
Equation 9. Here, p₁ is the pressure displayed by the integrity test unit (printed on the test report).

Effect of Pressure Reading Slope for Nonparallel Pressure Reading Offset: The diffusion value of the integrity test unit used for these trials is calculated from the pressure drop measurement. The correlation between the pressure drop measurement and the diffusion value is explained in Appendix B of PDA Technical Report 26 (1) and is not the objective of this article.

The integrity-test unit in those trials uses an improved algorithm according to DIN 58356, Part 2, taking into account the diminishing driving force as the pressure decreases over time as in
Equation 10. In that equation, D is diffusion in mL/min, p₁ is absolute starting pressure, V_net is the gas net volume of the filter setup; p₀ is reference pressure (typically set to 1,000 mbar), t is test time in minutes, and ▵p is the pressure drop during the measurement phase (▵p = p₁ – p₂).

To obtain an accurate estimation of the pressure reading offset influence, this logarithmic algorithm has to be taken into account using the actual starting pressure (p₁ – offset) as calculated above and the corrected pressure drop value we would have had if there had been no pressure reading offset (▵p_corr)

Calculation of the Corrected Pressure Drop: The slope m of the pressure reading offset curve might be different from the slope of the reference (1). In such cases, that difference will affect the measured pressure drop (Figure 8).

The pressure drop measured by the integrity-test unit and printed on the test report is the ▵p_read value. The pressure drop value that would have been measured if there had been no pressure reading offset ▵p_corr can be calculated using
Equation 11. Here, p_1x is the displayed starting pressure (printed on the test report) and p_2x is the pressure at the end of the test (p₁ – ▵p). The corrected pressure drop must be used in the equation. You can make an estimate by comparing the obtained result as measured by the integrity-test unit with the result obtained with corrected parameters.

The equation for calculating the impact of the difference in slope would then be
Equation 12, with with D_corr and D_printout defined in Equations
13 14 15 and simplified as shown in
Equations 16 and
17.

Clearly, the pressure reading offset will influence the diffusion value by

• the difference in applied test pressure resulting in a different driving force (as defined by Fick’s law)

• the eventual difference in pressure slope reading.

After calculating the correct initial test pressure and the corrected pressure drop value, we used them to estimate the effect on diffusion value, which was measured and printed by an integrity-test unit.

We calculated the expected deviation for a bubble point test with an instrument having a 200-mbar offset at full scale (compared with a calibrated instrument). Using the calibration values in Table 1, we achieved
Equation 18.

These trials show a very good correlation between theoretical and experimental values. An Excel spread sheet (Figure 9) for calculating the effects from an eventual offset was developed based on algorithms explained herein. For the case of a calibration offset, the calibration values can be entered into the spreadsheet and the overestimation or underestimation of the integrity-test values can be immediately evaluated based on a specific test pressure and surrounding calibration points.

Evaluating the impact as a percentage provides a fast and reliable risk analysis and can be comparable with the documented accuracy of the instrument during operational qualification (OQ). For example, if an instrument shows a diffusion reading error of +1% during OQ when it was perfectly calibrated and –3% based on the pressure reading offset evaluation with the spreadsheet, then its accuracy would still be within instrument specifications (±5%). Quality assurance personnel may then decide whether they need to check individual test results or if the deviation may be classified as without consequences.

The developed spreadsheet may or m
ay not be used for evaluating calibration offset on integrity test units from manufacturers than the one specified in this article.

Author Details
Magnus Stering is product manager for Integrity Testing Solutions, Sartorius Stedim Biotech; magnus.stering@sartorius-stedim.com.

REFERENCES

The post Effects of Pressure Sensor Calibration Offset on Filter Integrity Test Values appeared first on BioProcess International.

The biopharmaceutical industry is targeting high-concentration protein formulations to enable subcutaneous administrations. Such administration can provide better patient convenience than intravenous administration. One challenge associated with high-concentration formulations is increased electrostatic interaction between proteins and excipients. That is a result of increased protein-charge density at high protein concentrations. Such interactions can create an offset between excipient levels in final products and diafiltration buffers in ultrafiltration processes. The effect of such electrostatic interactions in a membrane process is known as the Donnan effect.

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Figure 1:  ()

The Donnan effect on excipient levels has received significant attention in recent years. Theoretical modeling has been developed to predict excipient and pH changes as a result of the Donnan effect in monoclonal antibody (MAb) processes. One model based on the Poisson–Boltzmann equation provided good prediction of excipient levels in the final retentate pool (1). A second model developed by Bolton et al. demonstrated to be predictive for basic MAb and acidic Fc-fusion proteins (2). The latter study also included several mitigation strategies to achieve target levels of excipients at the end of an ultrafiltration–diafiltration (UFDF) process. Both publications provide tools for understanding the influence of the Donnan effect on target formulation excipients. By contrast, our study focuses on the influence of the Donnan effect on removal of starting buffer excipients during diafiltration.

PRODUCT FOCUS: HIGH-CONCENTRATION BIOLOGICS
PROCESS FOCUS: FORMULATION, FILL AND FINISH
WHO SHOULD READ: PROCESS DEVELOPMENT AND MANUFACTURING, FORMULATION, FILTRATION OPERATORS
KEYWORDS: EXCIPIENTS, AGGLOMERATION, BUFFER REMOVAL, DIAFILTRATION
LEVEL: INTERMEDIATE

A typical final-formulation UFDF step will target eight to 10 diavolumes. For an ideal process, in which excipients pass freely through the membrane (the retention value R = 0), 10 diavolumes provide 99.995% removal of starting excipients. That equates to a “complete” exchange (Figure 1).

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Figure 1:  ()

The Donnan effect, however, can influence that removal. We performed several test runs to demonstrate how the Donnan effect changes the removal efficiency of positively and negatively charged excipients. We conducted diafiltration test runs using a MAb at two different concentrations to additionally assess the influence of MAb concentration on excipient removal efficiency.

Protein: We used SAN-300, a MAb provided by Santarus Inc., for the diafiltration studies. It is a glycosylated IgG1 monoclonal antibody directed against VLA1 (very late antigen-1, α1β1 integrin). The protein is expressed by a Chinese hamster ovary (CHO) cell line and purified using a standard three-column MAb purification process. SAN-300 protein has pI >8 with a molecular weight >140 kDa.

Excipients: We studied three different excipients in subsequent experiments. Two were negatively charged organic-acid buffers (referred to here as E_A^– and E_B^–). The third excipient was positively charged (referred to here as E⁺).

UFDF Procedure: We perfomed all tests at room temperature (21 °C) using EMD Millipore Ultracel (regenerated cellulose) 30-kD membranes in 88-cm² Pellicon 3 devices installed in a Pellicon Mini cassette holder. We ran diafiltrations at a transmembrane pressure (TMP) of 20 psig. The retentate was continuously stirred and recirculated through the system using a peristaltic pump. We performed initial concentration steps as needed to achieve the desired SAN-300 test concentration. Feed flow rate was 4.5 L/min/m² for the runs. Table 1 provides a summary of the test matrix.

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Table 1:&#8 194; ()

Measurement of Protein Concentration: Spectrophotometric methods determined protein concentration using the extinction coefficient at 280 nm. We diluted the reference standard and sample to a specified concentration and then measured at A₂₈₀, A₃₂₀, and A₃₆₀. We corrected the A₂₈₀ reading for background absorbance at A₃₂₀ and A₃₆₀. We then calculated protein concentration using the corrected A₂₈₀ dilution factor and extinction coefficient.

Excipient Assays: The positively charged excipient (E⁺) levels were determined using a capillary zone electrophoresis (CZE) method that uses a fused silica capillary (ID = 50 µm) with an enhanced detection cell, a borate electrolyte, and direct ultraviolet (UV) detection at 195 nm. We diluted the samples one hundredfold before analysis.

We measured the negatively charged excipients E_A^– and E_B^– using a special electrolyte for indirect UV detection of excipients, including a modifier to remove electroosmotic flow. We used a fused silica capillary (ID = 75 µm) and performed indirect detection at 200 nm. For both methods, we calculated concentration levels using the standard addition method.

RValue: We calculated the apparent retention of the excipients using
Equation 1 (see Equations box). N is the number of diavolumes.

We tested three starting/diafiltration excipient combinations for removal efficiency at 55 g/L and 120 g/L MAb concentration values. A total of six diafiltration runs were performed.

The first combination consisted
of a positively charged MAb, a negatively charged starting excipient (E_A^–), and a negatively charged diafiltration excipient E_B^–. Figure 2 shows the results as Runs 1 and 3. In that figure, the black line plot shows removal efficiency for a system in which excipients pass freely through the membrane with no effect from charge interactions (99.995% removal in 10 diavolumes). Removal efficiencies for E_A^– at 55g/L (Run 1) and 120 g/L (Run 3) were comparable to solutes with apparent retention values of R_app = 0.419 and R_app = 0.770, respectively. As Table 2 shows, when those results are compared with an excipient that passes freely through the membrane, seven additional diavolumes at 55 g/L protein concentration or 33 additional diavolumes at 120 g/L protein concentration would be needed to achieve a 99.995% removal.

Equation 1

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Equation 2

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Equation 3

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Equation 4

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Equation 5

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Figure 2:  ()

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Table 2:&#8 194; ()

We obtained similar results from the second combination, which consisted of a positively charged MAb, a negatively charged starting excipient E_A^–, and a positively charged diafiltration excipient E⁺, which corresponds to Figure 2, Runs 2 and 4. The negatively charged starting excipient E_A^– behaved as a partially retained solute with an apparent retention of R_app = 0.292 at the 55-g/L test condition. At the 120-g/L test condition, apparent retention increased to R_app = 0.419. For that case, a 99.995% removal would require 14 diavolumes at 55-g/L protein concentration or 23 diavolumes at 120-g/L protein concentration — compared with 10 diavolumes for an excipient that passes freely through the membrane (Table 3).

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Table 3:&#8 194; ()

The third combination consisted of a positively charged MAb, a positively charged starting excipient E⁺, and a negatively charged diafiltration excipient E_B^–. Starting excipient and MAb are both positively charged (Figure 2, Run 5 and Run 6). By contrast with the previous test results, positively charged E⁺ excipient exhibited enhanced removal efficiency. It provided 99.995% removal at <10 diavolumes. Removal efficiency was further enhanced at 120 g/L concentration (Table 4).

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Table 4:&#8 194; ()

Results showed that the removal efficiency of charged excipients during a diafiltration step was influenced by the Donnan effect. Excipients that had a charge that was opposite of the charge of the protein experienced electrostatic attractive forces, thereby partially retaining the excipient. That was demonstrated by the negatively charged excipient E_A^– and the positively charged MAb protein. By contrast, excipients with the same charge as the protein (e.g., positive– positive) experienced repulsive forces. For those cases, the excipient experienced enhanced removal comparable with a solute that passes freely through a membrane. Tests with E+ excipient and positively charged MAb demonstrated that result.

Protein concentration also influenced removal efficiency. At higher co
ncentrations, protein charge density increased, which in turn increased electrostatic interactions between protein and excipient (increased attractive or repulsive forces). That occurred with the diafiltration of E_A^– excipient using E_B^– diafiltration excipient. Diafiltering at 120 g/L protein concentration required 43 diavolumes, whereas 17 diavolumes at 55 g/L protein concentration provided a 99.995% removal, a 2.5× difference.

Because one objective of a final UFDF step is to provide a buffer exchange, process development scientists often target eight to 10 diavolumes under the assumption that process excipients pass freely through the membrane. Furthermore, optimum diafiltration concentration is typically based on hydraulic considerations (e.g., C_optimum = C_gel/e) (3).

Our data demonstrate that the number of diavolumes required to achieve excipient removal varies as a function of electrostatic interactions with diafiltration concentration. For excipients with a charge opposite to that of the protein, a diafiltration concentration below the C_gel/e optimum provided improved removal and lowered overall membrane area requirement.

To illustrate, we calculated membrane area requirements for diafiltration of the E_A^–/ E⁺ excipient combination (E_A^– removal using E⁺ diafiltration excipient) (Table 3). Calculations were based on a starting volume of 100 L at 55 g/L MAb, a diafiltration time of three hours, and a 99.966% E_A^– removal (Table 5).

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Table 5:&#8 194; ()

Table 5 shows that by assuming that E_A^– passes freely through the membrane, diafiltration at 120 g/L would require 51% less membrane area than needed for diafiltration at 55 g/L. By our analysis, a 120 g/L concentration would be a more optimal diafiltration concentration. If apparent retention due to the Donnan effect is considered, however, the number of diavolumes required to achieve 99.966% removal would be 11.3 and 18.4 for the 55 g/L and 120 g/L MAb concentrations, respectively. In this case, diafiltration at 55 g/L would require 39% less membrane area than would diafiltration at 120 g/L to achieve 99.966% removal. By our analysis, a 55 g/L concentration would be the optimum diafiltration concentration.

As described previously, the optimum concentration for diafiltration is generally determined from
Equation 2, in which C_gel/e represents the concentration at which the process area × time (m²h) value is minimized. This dependence of the optimum concentration on the C_gel value is determined on the assumption that excipients pass freely through the membrane (R = 0).

But because process area × time proportionality is as shown in
Equation 3 (in which R is the excipient retention and C_b is the concentration of product in the bulk solution), an expression for optimum diafiltration concentration that includes excipient retention can be determined. For example, if excipient retention is proportional to electrostatic forces (e.g., protein concentration) as suggested by our data, then
Equation 4 is reached (in which d is a proportionality constant).

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Figure 3:  ()

Equation 5 can be used to determine optimum concentration, in which optimum concentration for diafiltration is the C_b value that satisfies the equation. Figure 3 shows the left side and right side of the optimum concentration equation for the diafiltration process described in Table 3. For the E_A^–/E⁺ buffer combination, we determined the d value from the C_b and apparent retention values shown in Table 3. For this buffer combination with C_gel = 307 g/L and d = 0.005, we calculated C_optimum = 57 g/L. This example shows how an expression for excipient retention as a function of C_b can be used to determine optimum concentration for diafiltration that considers both hydraulic performance and electrostatic interactions.

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Figure 3:  ()

The Donnan effect influences removal of charged excipients during diafiltration of a high-concentration protein solution. Removal efficiency will be enhanced for excipients with the same charge as the protein and decreased for excipients with a charge opposite the protein. Removal of oppositely charged excipients is analogous to that of partially retained solutes.

Process development scientists should consider excipient removal when designing diafiltration steps. The required number of diavolumes to achieve excipient removal will depend on the extent of the Donnan effect, which is a function of protein concentration, protein charge, and excipient charge.

In certain cases, achieving a 99.995% removal may be impractical (e.g., high filter-area requirements). For such cases, process development scientists should identify acceptable removal targets. Proper development of the UF step can minimize process time and area while taking into account initial excipient removal, final excipient concentrations, and final protein concentration.

Author Details
Alexandra Steele and corresponding author Joshua Arias are bioprocess engineers at the Biomanufacturing Sciences Network unit of EMD Millipore Corp., 900 Middlesex Turnpike Rd., Billerica, MA 01821;1-781-533-2459; joshua.arias@emdmillipore.com.

REFERENCES

The post Accounting for the Donnan Effect in Diafiltration Optimization for High-Concentration UFDF Applications appeared first on BioProcess International.

Commercial-scale viral vaccine manufacturing requires production of large quantities of virus as an antigenic source. To deliver those quantities, a number of systems are used for viral replication based on mammalian, avian, or insect cells. To overcome the inherent limitations in production outputs with serial propagation of cells, mammalian cells can be immortalized, which increases the number of times they can divide in culture. Modifications that immortalize cells are typically accomplished through mechanisms similar to those converting normal cells to cancer cells. Thus, the presence of residual host-cell nucleic acids in final vaccine products would create significant concerns about the potential for transfer and integration into a patient’s genetic material.

PRODUCT FOCUS: VACCINES
PROCESS FOCUS: DOWNSTREAM PROCESSING
WHO SHOULD READ: PROCESS ENGINEERS, QA/QC, ANALYTICAL
KEYWORDS: CHROMATOGRAPHY, BIOCATALYSIS, TANGENTIAL-FLOW FILTRATION, NUCLEIC ACID DETECTION ASSAYS
LEVEL: INTERMEDIATE

Host-cell nucleic acids in feed material depends on cell/virus type and on methods and techniques used in harvesting. The presence of DNA can contribute to process fluid viscosity and fouling of separation media, reduce useable capacity, cause coprecipitation, and threaten product safety. The risk of oncogenicity and infectivity of host-cell nucleic acid can be minimized by suppressing its biological activity. That can be achieved by decreasing the amount of residual DNA and RNA and reducing their size (with enzymatic/nuclease or chemical treatment) to below the functional gene length of ~100 base pairs.

Precipitation with

• Cationic detergents — e.g., cetyltrimethyl ammonium bromide (CTAB) or domiphen bromide (DB)

• Short-chain fatty acids (e.g., caprylic acid)

• Charged polymers — e.g., polyethyleneimine (PEI) and polyacrylic acid (PAA)

• Polyethylene glycol (PEG)

• Ammonium sulfate

• Tri(n-butyl)phosphate (TNBP) with Triton X-100 detergent solution

Filtration:

• Normal-flow filtration (NFF) with depth-charged or diatomaceous-earth–containing media

• Tangential-flow filtration (TFF)

• Ultrafiltration/diafiltration (UF/DF)

Chromatography and membrane adsorbers:

• Anion-exchange chromatography (AEX)

• Gel-filtration (size-exclusion) chromatography

• Hydrophobic charge-induction chromatography (HCIC)

Degradation with

• Enzymes

• Physical forces (shearing)

• Alkylating agents (e.g., β-propylactone)

Health authorities and regulatory bodies such as the US Food and Drug Administration (FDA) and the European Medicines Agency (EMA) have set limits for acceptable amounts of residual DNA in final biological products. According to requirements published by the FDA, a parentally administered dose is limited to 100 pg of residual DNA. The EMA and the World Health Organization (WHO) allow 10 ng per parenteral dose and 100 μg/dose for an orally administered vaccine (1). Orally administered DNA is taken up about 10,000× less efficiently than parenterally administered nucleic acid.

The biopharmaceutical industry continues to improve its purification processes to minimize potential risks associated with harmful immunological and biological responses caused by residual impurities originating from host cells and culture media. Here we describe a new method for removal of host-cell DNA and/or RNA impurities that offers some advantages over known approaches. We also summarize our development of a process incorporating this new approach.

A typical cell-culture–based viral vaccine production process begins with propagation of a selected “seed” cell line. When the culture has grown to a predetermined cell density, the virus of interest is introduced (inoculated) and begins to replicate. A few days later, virus is harvested directly from the host cells, from the culture supernatant, or both.

When virus is harvested from the host cells directly, a cell disruption step is required to release intracellular viruses. Available methods for cell disruption fall into two main categories: chemical (detergents and surfactants, enzymes, osmotic shock) and physicomechanical (heat, shear, agitation, sonication, freeze-thawing). Mammalian cells are easy to break, so the methods used for their disruption are relatively mild and may include treatments like use of detergents and/or hypotonic buffer.

When virus is harvested from supernatant, there is no need for a cell breakage step; the supernatant can be directly clarified from cellular debris. If a virus must be harvested from both cells and supernatant, collected cells will be disrupted before the viral suspension is clarified.

The amount of host-cell–related impurities (including nucleic acids) in a process fluid varies significantly depending on the methods used for cell lysis and/or virus harvest. The “DNA Removal” box lists several techniques that can be applied for reduction and/or removal of genomic DNA from cell culture process streams. But all of those methods are limited in the types of products and processes for which they could be applied.

Purification of viruses from cell substrate components such as DNA is a particularly challenging task in viral vaccine production for a number of reasons:

• Physical similarities of viruses and nucleic acids could limit the resolution and selectivity of an applied method. For example, similar electrical charge (both viruses and nucleic acids are typically negatively charged at neutral pH) and size create limitations in separation with chromatography and filtration methods (Table 1). And sedimentation behavior similarities could lead to coprecipitation.

• Applied techniques and reagents could affect biological virus activity and integrity, causing losses of infectivity and/or potency due to degradation by physical forces (shear) or chemicals (detergents). Caprylic acid, for example, can inactivate enveloped viruses.

• Some methods can cause nucleic acids to bind with a product of interest (virus/glycoprotein/protein), which lessens the efficiency of downstream purification processes in achieving the necessary level of final-product purity and compromising process yields and economics.

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Table 1:&#8 194; ()

Most of those limitations and challenges can be minimized when the amount of nucleic acid contaminant is reduced using enzymatic degradation with endonucleases. Such enzymes act by specifically catalyzing the hydrolysis of the internal phosphodiester bonds in DNA and RNA chains, breaking the nucleic acids into smaller nucleotides. Naturally present in bacteria, these enzymes defend cells from invasion by foreign DNA. The several types of these nucleases are derived from different sources.

Serratia marcescens is a Gram-ne
gative pathogenic bacterium that secretes (among other proteins) a very active endonuclease that cleaves all forms of DNA and RNA (single-stranded, double-stranded, linear, and circular) without sequence specificity. The nuclease cleaves nucleic acids very rapidly, with a catalytic rate almost 15× faster than that of deoxyribonuclease I (DNase I) (2). The enzyme shows long-term stability at room temperature and is active in the presence of both ionic and nonionic detergents as well as many reducing and chaotropic agents. But it has proteolytic activity of its own. All these characteristics make this enzyme useful for biotechnological and pharmaceutical applications.

A genetically engineered form of Serratia nuclease (Benzonase from Merck Millipore of Darmstadt, Germany) is made using Escherichia coli and recombinant technology. The Benzonase enzyme is a dimer of identical subunits with molecular weight ~30 kDa each (with a weight totaling ~60 kDa). Its isoelectric point is at pH 6.85, and it is functional in a pH range of 6–10 and at temperatures of 0–42 °C. The presence of Mg2⁺ (at 1–2 mM concentration) is required for this enzyme’s activity.

The Benzonase enzyme digests all forms of nucleic acid by hydrolyzing them into smaller oligonucleotides of <10 base pairs in length (1 bp is ~650–660 Da) with minimal sequence specificity. (Hydrolysis occurs slightly more often in guanosine and cytosine (GC)–rich areas than in adenine and thymine (AT)–rich sequences). During a typical reaction, nucleic acid molecular weights are reduced as in Figure 1 through ethidium-bromide–stained agarose electrophoresis gels.

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Figure 1:  ()

One Benzonase unit is defined as the amount of enzyme that causes a change in UV absorbance at 260 nm of one absorption unit within 30 minutes. That unit degrades about 37 μg DNA in 30 minutes.

Because process times for enzymatic reactions often measure in hours, Benzonase treatment is typically carried out in batch mode. To start a reaction, the enzyme is added to a process feed. Large amounts are often used to ensure maximal digestion of nucleic acid impurities. In such cases, no optimization of the enzymatic reaction is needed; process step optimization takes into account temperature, exposure time, and shear on product stability rather than on specific Benzonase activity.

Current examples for laboratory scale applications demonstrate use of 9–90 U/mL. If required, the enzymatic reaction can be optimized through studying reaction rates in microwell plates because only very small quantities are needed both for reaction and DNA/RNA detection. After treatment, subsequent purification steps must quantitatively remove the enzyme from a process stream. Benzonase treatment therefore should be placed sufficiently upstream in the overall process (Figure 2).

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Figure 2:  ()

Critical parameters of the enzymatic reaction to consider during process development include enzymatic activity, DNA and RNA feed and target concentrations, process time, enzyme and Mg2⁺ concentrations, temperature, pH, and the presence (and concentration) of Benzonase inhibitors and multivalent or monovalent salts in media and their concentrations. And typical enzymatic reactions can be described by Michaelis– Menten kinetics (
Equation 1).

The volumetric rate of reaction (vrr) is proportional depending on the maximum reaction rate at infinite reactant concentration (vrr_max) and on the substrate concentration (S) of nucleic acids DNA and RNA and the Michaelis constant (Km).

The highest Km is achieved by operating at optimal pH (8–9) and temperature (37 °C). Although higher temperatures expedite the reaction kinetics, the need for simplified control and/or concerns over product stability often lead to operating at room temperature or lower. Mg2⁺ ions are needed to get optimal enzymatic activity.

Another critical parameter is the enzyme’s starting activity, so enzyme concentration is provided in units (as described above) rather than actual concentrations. When concentration of host-cell nucleic acids in starting feed material is high, the maximum reaction rate is independent of the their concentration. As enzymatic digestion progresses and nucleic acid concentration is reduced, the reaction will become first order and the removal rate is reduced.

Some process additives and agents affect Benzonase activity. The enzyme can be inhibited by high salt concentrations (Figure 3): >300 mM monovalent cations, >100 mM phosphate, >100 mM ammonium sulfate, or >100 mM guanidine HCl. Other known inhibitors include chelating agents. EDTA, for example, could cause loss of free Mg2⁺ ions (EDTA concentrations >1 mM have shown to inhibit the enzymatic reaction), an effect that can be reversed by adding more MgCl₂. And the presence of components such as 4M urea could have an opposite effect and increase Benzonase activity.

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Figure 3:  ()

Because the starting material might contain specific inhibitors, concentrations, or salt choices — and enzyme quality (in terms of activity) is predefined — process optimization at a given position in the process can focus on controllable variables as a function of process time: enzyme and Mg2⁺ concentration, temperature, pH, and salt co
ncentrations. The information herein can be used for troubleshooting purposes for cases in which excessive amounts of enzyme are used.

Regulatory authorities do not regulate how much residual endonuclease can be in a vaccine product. However, vaccine manufacturers using it in their processes need data to demonstrate safety/toxicity status and measure residual endonuclease that might be present in final preparations. Consider Merck & Company’s EU patent of VAQTA hepatitis A vaccine (3). It indicates that residual Benzonase enzyme is lower than 0.0001 ng/dose. It is important to note that the endonuclease is a process additive and not a drug, excipient, or active pharmaceutical ingredient.

Benzonase removal from a vaccine process stream can be accomplished by several downstream unit operations, so Benzonase treatment is often positioned in the “upstream” part of processing. Removal can be demonstrated by showing a lack of residual nuclease activity (which does not detect residual nonactive enzyme) and using an enzyme-linked immunosorbent assay (ELISA) for detection of total residual Benzonase molecules (both active and nonactive).

Irreversible Benzonase inactivation occurs within ~15 min at a temperature >70 °C and 0.02 N NaOH. Such conditions could negatively affect the integrity of viral vaccines, however. Heating the product solution also could increase Benzonase activity, thus affecting the determination of residual nucleic acids in samples. Removal of the enzyme can be accomplished using classical downstream methods, as described below. A clearance technique can be chosen to align with additional purification steps for the vaccine.

Tangential-Flow Filtration (TFF): Benzonase enzyme is removed in filtrate while viral particles are retained. For this ~60-kDa molecule, we suggest 300-kDa Biomax or larger cut-off sizes. Depending on the viral particle size (viruses should be retained with low passage), TFF might be a possible option. An internal study at our company indicated that Benzonase enzyme could be removed by diafiltration ≤99.5% at five diavolumes and >99.9% after eight diavolumes using 300-kD membrane. The overall diafiltration profile is close to a theoretical sieving value of 1. We recommend 300 kD as an acceptable molecular-weight cut-off (MWCO) value for the Benzonase clearance after DNA digestion in viral cell cultures — provided that the membrane is retentive enough for the virus of interest.

Size/amount estimation of Benzonase spike required for treatment of cell substrate for gDNA digestion for a live/attenuated virus for injectable vaccine product

Virus propagation system: adherent VERO cell culture

Cell lysis type: chemically induced

Product stream: postsecondary clarification (NFF) viral suspension

Host cell/gDNA content: ~1 μg/mL

Batch volume (product stream of interest): ~10 L

Total amount of gDNA: 10,000 μg

Minimum Benzonase amount for gDNA digestion: ~270 U (0.027 U/mL)

Benzonase type to apply: HC, >99% pure, 25 KU, Catalog#71206-3 (250 U/μL)

Benzonase amount/spike applied: 500 U (=2 uL)

Typical amount used: 20 U/mL (minimum) to 50 U/mL (maximum)

Safety factor used: 740 (minimum) to 1,852 (maximum)

Anion-Exchange Chromatography (AEX) is considered to be the most convenient and effective chromatography technique for Benzonase removal. With a pI of 6.85, the enzyme will typically flow through while a viral product is bound to an AEX column or (if it is bound as well) elutes separately. Table 2 lists several applicable AEX resins using a number of sample and equilibration buffers.

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Table 2:&#8 194; ()

Cation-Exchange Chromatography (CEX) has been shown to remove Benzonase enzyme, but the operating range might be smaller than for AEX. Table 3 lists a few CEX chromatography media and conditions that are suitable for the Benzonase removal.

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Table 3:&#8 194; ()

As chemical composition, process time, and process temperature are critical parameters, it is important during scale-up to ensure that the entire solution is mixed efficiently so that all parts of a batch experience similar enzymatic reaction conditions. That can be achieved using well-characterized mixers and thermal control. Understanding mixing and temperature equilibration is critical in reaction scale-up. Take care to ensure that treated material is not contaminated with untreated material (e.g., residual material in a transfer line or splashed material that is not part of the batch reaction). Some viruses are shear sensitive, so confirmation of viral product stability during mixing needs to be verified. Mixing speed thus might be explored as an area of focus. The “Case Study” box above provides a typical example of Benzonase use sizing.

The residual DNA limit of 100 pg/dose set by regulatory authorities equals the DNA amount from ~17 diploid Chinese hamster ovary (CHO) cells (4). Detection of such a small amount of DNA requires an extremely sensitive and robust analytical method. The European Pharmacopoeia advises that residual DNA should be determined using sequence-independent techniques (5). Common methods of measuring DNA include UV absorbance at 260 nm (maximal absorbance with an extinction coefficient of 50), fluorometric detection using ethidium bromide (EtBr) or Hoechst 33258, a Picogreen assay (Life Technologies), quantitative polymerase chain reaction (qPCR), and a Threshold DNA assay (Molecular Devices). Table 4 summarizes their detection limits.

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Table 4:&#8 194; ()

Components in the feed mixture may affect assay results. To assess the level of digested DNA bands, ultralow-range DNA ladders — ≥10 bp according to polyacrylamide gel electrophoresis (PAGE) or agarose-electrophoresis measurements — could be used (6)(7). Before qPCR analysis is performed, Benzonase enzyme must be inactivated or removed; otherwise, it may digest newly amplified DNA. Adding an ice-cold solution of perchloric acid (4%) will instantly stop the Benzonase activity. The Threshold DNA assay is not based on hybridization, but rather relies on reaction chemistries and luminescence. Reported detection limits are as low as 4 pg/mL (2 pg for a 0.5-mL sample). DNA log reduction factors (LogRF) can be calculated using
Equation 2.

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Several methods are used for removing nucleic acids from bioprocess streams, all of which are still valid for current processes. But new cell lines pose challenges to drug safety. It is important to eliminate host cell DNA/RNA impurities by thoroughly removing them from a product stream. Benzonase enzyme offers some advantages over competing methods of nucleic acid removal. Not designed to be part of the final product, however, it needs to be removed after use.

Benzonase removal from a vaccine process stream can be accomplished using several types of downstream unit operations. Depth filtration for clarification, TFF for concentration and diafiltration, and chromatography for purification can also remove nucleic acids. The latter two could be used to ensure Benzonase removal from treated product streams. Results can be demonstrated by a lack of residual nuclease activity and with an ELISA for detecting total residual endonuclease.

Author Details
Elina Gousseinov, MS, is a process development scientist at EMD Millipore in Canada. Willem Kools, PhD, is head of field marketing and biomanufacturing sciences at EMD Millipore in Bedford, MA. And corresponding author Priyabrata Pattnaik, PhD, is director of the worldwide vaccine initiative at Merck Millipore, 1 Science Park Road, #02-10/11 The Capricorn, Singapore 117528; 65-6403-5308; priyabrata.pattnaik@merckgroup.com. Benzonase is a registered trademark of Merck Millipore.

REFERENCES

The post Nucleic Acid Impurity Reduction in Viral Vaccine Manufacturing appeared first on BioProcess International.

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Figure 1: Filtration principle of dynamic body-feed filtration (DBF) with diatomaceous earth (DE) (left) and conventional filtration (right)

In the past decade, biopharmaceutical manufacturers have demonstrated major improvements in monoclonal antibody (MAb) production, exhibiting product titers frequently in the range of 5–10 g/L using standard fed-batch mammalian cell cultures (1, 2). Increased product yields allow for smaller-scale production vessels. With 2,000-L single-use bioreactors already commercially available, single-use manufacturing of biomolecules becomes more and more an option. Other recent developments in the biopharmaceutical industry — e.g., drugs for smaller indications and more potent drugs allowing for lower dosages — will further stimulate the demand for smaller and more flexible single-use manufacturing facilities.

Although single-use technology in general has matured considerably over the past few years, some unit operations (e.g., cell removal) still need more attention to become more economical and robust. High product titers often result from increased cell densities rather than increased specific productivities per cell, and the resulting solids content poses considerable challenges on commonly applied harvesting technologies. Currently the most prevalent single-use harvesting technology, depth filters block at lower loading capacities with higher biomass concentrations. Higher contaminant concentrations also make depth filters more sensitive to batch variation, which can lead to 50% oversizing of filter area to compensate for fluctuating filtration capacities. That drives up costs and increases waste. Other new commercially available single-use cell-removal technologies such as centrifuges still lack capacity. For harvesting higher–cell-density cultures, a major technical breakthrough would be welcomed.

Body Feed Filtration Successful for Plasma Fractionation

When we looked at other industries that have similar process needs — such as the plasma fractionation industry — we found that they often use body-feed filtration for clarification of solutions with high solids content. The first use of diatomaceous earth (DE) as a filter aid in fractionation of human plasma was reported over five decades ago (3). Since then, a number of manufacturing processes for production of intravenous immunoglobulin (IVIG), albumin, and clotting factors have been developed based on that technology (4–6). Fractionation uses the principles of selective precipitation by pH adjustment, ionic strength, addition of alcohol, and temperature shifts. Precipitates are removed by depth filtration often in combination with diatomaceous earth, which is added as an aid to increase filter throughputs.

Recently, the principles of body-feed filtration have been tested for harvesting cell cultures, with promising results (7). Our objective was to evaluate the technology as a potential single-use alternative to centrifuges and depth filters. Here we describe the most interesting findings we obtained using a number of cell lines and culture media, which were kindly provided by different biotech companies. Together with Rentschler Biotechnologie GmbH, Sartorius Stedim Biotech tested the optimized conditions in a 600-L cell culture production process to evaluate the scalability of body-feed filtration technology.

How Body Feed Filtration Works

When the concentration of solids or colloids is too high in turbid biological process fluids needing clarification, the filter cake forming on the surface of a filter becomes impermeable and blocks the filter (Figure 1, right). Adding highly porous DE creates a more permeable filter cake, which prevents blockage (Figure 1, left).

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Figure 2: Mean particle-size distribution of cell-free Chinese hamster ovary (CHO) culture
supernatants from three different harvest days determined at different pH values; green curve = pH 7.0, red curve = pH 5.0; measurement performed using a Mastersizer 2000 system (Malvern Instruments)

The minimum amount of DE to guarantee smooth filtration depends on the particle concentration. Laboratory-scale experiments with many different mammalian cell lines, culture media, and viabilities revealed a correlation between the wet cell weight (WCW) and the required amount of DE at constant pH (data not shown). For all tested cultures, the optimum DE concentration was in the range of 40–50% of WCW. In all cases, the filter-aid ratio could be reduced significantly to a range of 20–30% when pH was lowered to pH 5.

Low pH Precipitation Improves Clarification Results

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Photo 1: Scanning electron microscopy (SEM)
shows the porous structure of Celpure 300 diatomaceous earth (magnitude 1,000 times).

Performance differences between acidified (pH 4.3–5.5) and neutral cell culture fluids for other cell-removal technologies such as microfiltration (4) and depth filtration (5) can be explained by precipitation of smaller particles at lower pH levels. The solubility of cell debris and negatively charged impurities such as DNA and host-cell proteins (6) decrease along with pH. Figure 2 shows the mean particle-size distribution for three different cell-free cell culture supernatants at neutral pH (green line) and after the pH of those cultures was lowered to pH 5 (red line). Lowering the pH leads to formation of bigger particles and makes the typical submicron particles (<1 μm) present at pH 7.0 completely disappear. Therefore, body-feed filtration would clearly benefit from lowered pH.

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Figure 3: Turbidity measurement of the DBF filtrates at pH 7.0 and pH 5.0 for 10 different
cell cultivations with an initial turbidity of
2,396–3,235 NTU

In addition to the significantly reduced filter-aid concentration, all tested acidified cell culture supernatants showed much clearer filtrates than their neutral counterparts (Figure 3). When we analyzed the particle-size distribution of neutral body-feed filtrates, we detected only small particles (<0.4 μm). Filtration with a 0.2-μm membrane barely reduced those turbidity values, indicating that very small particles are mainly responsible for the higher turbidity values of the neutral body-feed filtrations.

The absence of smaller particles in acidified culture supernatants probably is also the reason for higher filtration capacities at constant filter-aid concentrations. We assume that small particles deposit in the pores of DE particles, lowering the permeability and filtration capacity of the DE overall. By increasing the filter-aid concentration, we could increase its capacity for retaining small particles without losing the required cake permeability.

Significant reduction of filter-aid consumption at low pH is promising from both economical and handling points of view. Another important economical factor to consider is the recovery rate at lower pH levels. Although most antibodies are stable at acidic conditions (9), in some cases low-pH precipitation of cell culture fluid will lower recovery rates, most likely due to coprecipitation of the antibody (6). However, some authors have described that lowering the pH value influenced overall antibody recovery positively by preventing enzymatic reduction of the product (10, 11). In all our laboratory-scale body-feed tests at low pH, we achieved a recovery rate >85% (data not shown).

Positive Impact on Residual DNA and Host-Cell Protein Load

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Figure 4: Mean residual DNA amount after DE filtration at pH 5.0 for 10 different cell culture supernatants, expressed in percentage of initial
DNA concentration at pH 7.0, measured in the cell free supernatant; initial concentrations
were 475–730 ppm (DNA quantitation with PicoGreen assay from Life Technologies).

With the low-pH precipitation and subsequent retention of contaminants by body-feed filtration, we accomplished a significant reduction of DNA in the primary recovery step from our cell culture runs. Figure 4 shows the mean residual DNA content of 10 different body-feed filtrates after low-pH precipitation at pH 5.0. We first measured the initial DNA content in the neutral, cell-free supernatant of each cell culture at pH 7.0. After a low-pH precipitation step followed by DE filtration, DNA content in the cell culture supernatants was reduced by 65%.

Pilot-Scale Test Results

At Rentschler Biotechnologie, we performed tests at pilot production scale to evaluate the applicability of DBF technology for manufacturing purposes (12). In total, 1,000 L of a high–cell-density (17.6 × 10⁶ cells/mL, 95% viability) Chinese hamster ovary (CHO) cell fed-batch culture were available for the depth-filtration and body-feed filtration runs. In the largest single run, we subjected 600 L to body-feed filtration at pH 5.0. WCW on that day was 8%.

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Figure 5: Schematic representation of pilot-scale body-feed filtration

For the scale-up experiment, we installed seven process-scale modules with a total filtration area of 1.61 m² in a universal stainless steel holder. The filter cassettes consist of two polyethylene filter plates, which retain the DE and biomass (Figure 6).

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Figure 6: (top) 3-D illustration of commercial
DBF process-scale module holder; (center) a drawing of one single-use DBF module; (bottom) flow path inside one module indicates in red the feed flow and in green the filtrate flow path; dotted lines indicate the filter plates.

In total, we added 12 kg of Cellpure C300 DE to the 600-L bulk harvest. Prefilled DE bags were connected with a dust-free adapter to a mixing bag for fast and safe DE transfer. Just five minutes of gentle mixing was sufficient to dissolve that DE powder in the cell suspension. Before filtration, we adjusted the pH of the resulting mixture to a final level of 5.0 and gently mixed it for two hours at 140 rpm.

Pressure increased steadily during filtration (Figure 7, left). We terminated filtration when the pressure reached 1.3 bar and the crude harvest had been filtered. During the entire filtration process, the system maintained a high and stable flux slightly above 300 L/m²/h. Overall, a capacity of 311 L/m² was achieved. We monitored a very low turbidity of 5–8 NTU (Figure 7, left) in the clarified harvest stream during filtration.

After neutralization, pool 3 (the final pool) exhibited a turbidity of 41 NTU (Figure 7, right), which was considerably higher than turbidity had been during filtration. Inadequate dosing of the neutralization buffer probably had resulted in local pH excursions and caused the turbidity increase in that final pool. In a small-scale parallel test, the turbidity increase was prevented through more gentle neutralization of the filtrate pool. To improve that neutralization step, an integrated ready-to-use process skid is under development that will enable controlled inline pH adjustment and prevent overshooting.

We found IgG1 recovery to be acceptably high at 85% (Figure 7, right). In the future, an optimized neutralization procedure and larger postfiltration flush should further improve MAb recovery. We monitored contaminants such as a host-cell protein and DNA throughout the process. In the final pool (after buffer flush and neutralization), those levels were reduced from 841 to 629 mg/mL and 13.8 to 5.0 μg/mL, respectively.

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Figure 7: Results of DE body-feed scale-up experiment at reduced pH (5.0) with seven filter modules; filtration performance (left), pressure (bar), and flux (L/m²) as well as the course of turbidity during filtration; recovery of IgG1, blue columns (right) measured from crude harvest and harvest pool without buffer flush (pool 1), with buffer flush (pool 2), and after neutralization of harvest fluid (pool 3)

Concluding Remarks

Our aim was to demonstrate the universal applicability of a new single-use harvest method for mammalian cell culture, suitable even for high–cell-density cultures. Tests using crude harvests from different cell lines and culture conditions allowed us to determine the optimal concentration of DE as a filter aid in relation to WCW, which is an easily accessible process specific for cell removal and harvest processing. With respect to process economics, the 50% reduction of filter aid required with low-pH filtrations is promising.

Generally, the pilot-scale results confirmed our findings from DBF filtration trials at laboratory scale: Reducing pH to 5.0 after addition of DE to crude cell-culture supernatant gives the best performance in terms of filtration capacity, flux, and contaminant removal.

The applicable flux rate of DBF technology is very advantageous. A 600-L harvest was processed within only an hour of filtration using just seven modules. A module holder allows arrangement of a maximum 33 modules, so we estimate that a harvest volume of ~3,000 L could be filtered in the same time.

In conclusion, this method allows effective clarification of high–cell-density, crude cell culture harvests in a single-use set up at large scale. Economically and competitively, it can replace centrifugation, which is currently the method of choice for large-scale cell removal. Even very dense crude cell harvests could be clarified quickly at high flow rates. Moreover, this method has the additional benefit of efficiently reducing contaminants in a single step. Scalability — one of the most important requirements in bioprocessing — is easily attainable following a generally linear approach with very consistent process performance.

References

1 Kelley B. Industrialization of MAb Production Technology: The Bioprocessing Industry at a Crossroads. MAbs 1(5) 2009: 443–452.

2 Buttler M. Animal Cell Cultures: Recent Achievements and Perspectives in the Production of Biopharmaceuticals. Appl. Microbiol. Biotechnol. 2005, 283–291.

3 Cohn EJ, et al. Preparation and Properties of Serum and Plasmaproteins, IV: A System for Separation into Fractions of the Protein and Lipoprotein Components of Biological Tissue and Fluids. J. Am. Chem. Soc. 68, 1946: 459–475.

4 Curling J, et al. Production of Plasma Proteins for Therapeutic Use. Wiley-Blackwell: Hoboken, NJ, 2012.

5 More J, et al. Purification of Albumin from Plasma. Blood Separation and Plasma Fractionation. Harris J, Ed. Wiley-Liss: Hoboken, NJ, 1991.

6 Stucki M, and al. Investigations of Prion and Virus Safety of a New Liquid IVIG Product. Biologicals 36, 2008: 239–247.

7 Westoby M, et al. Effects of Solution Environment on Mammalian Cell Fermentation Broth Properties. Biotechnol. Bioeng. 108(1) 2011: 50.

8 Delavaille D. How to Apply Old Techniques to New Processes. European Downstream Technology Forum, 7–8 September 2010, Göttingen Germany. Sartorius Stedim Biotech AG: Goettingen, Germany.

9 Singh N, et al. Clarification of Recombinant Proteins from High Cell Density Mammalian Cell Culture Systems Using New Improved Depth Filters. Biotechnol. Bioeng. 110(7) 2013: 1964.

10 Brodsky Y, et al. Caprylic Acid Precipitation Method for Impurity Reduction: An Alternative to Conventional Chromatography for Monoclonal Antibody Purification. Biotechnol. Bioeng. 109(10) 2012: 2589.

11 Kao Y, et al. Mechanism of Antibody Reduction in Cell Culture Production Processes. Biotechnol. Bioeng. 107(4) 2010: 622.

12 Trexler-Schmidt M, et al. Identification and Prevention of Antibody Disulfide Bond Reduction During Cell Culture Manufacturing. Biotechnol. Bioeng. 106(3) 2010: 452.

13 Minow B, et al. High-Cell-Density Clarification By Single-Use Diatomaceous Earth Filtration. BioProcess Int. 12(4) 2014: S36 –S47. c

Tjebbe van der Meer, MSc, is a product manager at Sartorius Stedim Biotech GmbH, August-Spindler-Straße 11, 37079, Göttingen, Germany, tjebbe.vandermeer@sartorius-stedim.com. Benjamin Minow, PhD, is director of cell culture disposable manufacturing at Rentschler Biotechnologie GmbH, Erwin Rentschler Straße 21, 88471 Laupheim, Germany, benjamin.minow@rentschler.de. Bertille Lagrange, MSc, is a scientist at Sartorius Stedim Biotech GmbH, August-Spindler-Straße 11, 37079, Göttingen, Germany, bertille.lagrange@sartorius-stedim.com. Franziska Krumbein, Dipl. Ing (FH), is director of process technologies at Sartorius Stedim Biotech GmbH, August- Spindler-Straße 11, 37079, Göttingen, Germany, franziska.krumbein@sartorius-stedim.com. And Francois Rolin, Dipl. Ing, is with ChangeXplorer Production SAS, Z.I. des Waillons 02220, Braine, France changexplorer@orange.fr.

The post Diatomaceous Earth Filtration: Innovative Single-Use Concepts for Clarification of High-Density Mammalian Cell Cultures appeared first on BioProcess International.

According to current European Union good manufacturing practice (EU GMP), integrity testing of sterilizing-grade product filters should be performed preuse poststerilization (PUPSIT) and immediately after use. In addition, PDA’s Technical Report 26 states that preuse integrity tests are preferably performed after filter sterilization. Performing an integrity test of an already sterilized product filter in-line requires wetting the filter while maintaining the downstream side sterile. The test gas must also be evacuted on the downstream side throughout testing maintaining sterility. The upstream side also must be protected from uncontrolled bioburden, which generally is understood as maintaining sterility of the upstream side by using sterile water for wetting and a sterilizing barrier between the integrity tester and the filter to be tested.

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Photo 1: The filter management system (FMS)

The drawback of performing integrity testing on a sterilized filter in-line is that rather than increasing process safety, it can increase risk from operator-handling errors and additional filters. The installation also becomes more complicated. The most common method of maintaining sterility on the downstream side throughout wetting and during a test is to use a sterilizing-grade filter, which has to be tested to prove maintained sterility. In addition, the supply of WFI from the upstream side typically requires a sterilizing-grade filter, which then has to be integrity tested to demonstrate maintained sterility.

Taking into account all filters required to protect a process filter and a process line for implementing preuse poststerilization integrity testing, at least three to four additional integrity tests must be performed. In addition to the increase of risk from these filters, there is significant down time of the installation.

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Photo 2: Housing with downstream valve

Here I present an engineering approach for preuse poststerilization integrity testing that already has been implemented on several production sites. Instead of blocking the process line, filter integrity is performed in a filter’s housing (in situ) on an automated test bench (off line). This in-situ off-line approach strongly reduces the downtime of a process line as a filter in its filter housing is being tested, while an already integrity tested identical filter in an identical filter housing are installed on the process line.

Filter Management System
Rather than installing a new filter to be used in its filter housing on a process line, the filter is installed in its filter housing on a test bench (Filter Management System, FMS) (Photo 1).

A fully automatic process is launched in the following steps:

• Steaming the filter in its housing (after preflushing with water, if required)
• Precooling the filter and housing under sterile conditions
• Wetting with sterile water and draining the filter under sterile conditions
• Integrity testing the filter under sterile conditions (with temperature survey if defined in the user requirement specifications)
• Drying the filter and its housing under sterile conditions • Closing upstream and downstream valves included on the housing (Photo 2, only downstream valve is shown).

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Figure 2: Downstream sterilization separate from the installation

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Figure 1: Downstream steam sterilization with the installation

The closed filter housing (including the integrity-tested sterilizing-grade filter) can then be installed on a process line. The downstream connection is steam sterilized either together with the installation (Figure 1) or separately before the valve is opened (Figure 2). The upstream connection also can be steam sterilized. Because every filter housing has got an identical “twin-brother housing,” one housing may be in use while the other is processed on the FMS with a new filter (or reused in case of a hydrophobic vent filter).

Because the filter is dried after the integrity test and kept sterile by the closed valves, it can be put on stock before being used in the process. Figure 3 shows the life cycle of a filter in its filter housing.

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Figure 3: Life cycle of a filter in its housing

Steaming a Filter in Its Housing: The automatic process on the FMS method starts with steaming a filter (preceded by a preflushing of the hydrophilic filter cartridge if required by the filter’s manufacturer). Steaming should be supervised in terms of temperature and time and data recorded in a database. Becasue no additional equipment is steam sterilized with the filter, steaming conditions are obtained with minimal differential pressure applied over the membrane. And because the process is supervised, there is no risk of being outside the filter’s temperature and differential-pressure specifications.

Precooling Filter and Housing under Sterile Conditions: To prevent stress on a filter cartridge (as from fast cooling when flushed immediately with water), the housing is cooled down gently with sterile air to nearly 60 °C. The integrated air filter on the FMS — which is used to supply sterile air — is automatically integrity tested before and after the process, thus ensuring that sterility is maintained.

Wetting and Draining a Filter under Sterile Conditions: After precooling, the filter cartridge can be cooled down by flushing with sterile room-temperature water. Sterile water is obtained by filtering water for injection (WFI) through an integrated sterilizing-grade filter. This filter is automatically integrity tested before and after the process, thereby ensuring that sterility is maintained.

Fully automated flushing implements a higher inlet pressure and back pressure than conventionally used when wetting filter cartridges before integrity testing. The back pressure is obtained by limiting the flow, and the differential pressure over the membrane is kept low for minimal stress on the membrane. This procedure provides more efficient wetting conditions, thus requiring a lower volume of water. Such wetting conditions contribute to process consistency because nearly 90% of all filter-failure claims from customers are a result of inadequate flushing.

If the filter to be tested is a hydrophobic filter, then the filter cartridge cannot be rinsed with water. The filter housing is therefore filled with sterile water to cool it down. The cooling water is then drained, and the filter housing is refilled to perform the water intrusion test (WIT) or water flow test (WFT).

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Figure 4: Steam-through connector

Filter Integrity Testing under Sterile Conditions: Integrity is tested according the pressure-drop method, for which the correlation to the integrity test value is explained in PDA Technical Report 26 (1). The filter housing including the filter to be tested is pressurized using sterile air. After a predefined stabilization time at the test pressure, the housing gas net volume is measured using a reference volume. The reproducibility of the net volume value confirms that the correct filter size has been installed in the housing.

After the net volume is measured, the test pressure is restabilized and the pressure drop is measured. During the test, a temperature survey is performed and recorded (if specified in the user requirement specifications) to monitor temperature influences. After the integrity test, a full test report is displayed and recorded in the data system. Printing can be performed locally and/or in a different area as required.

Drying the Filter and Its Housing under Sterile Conditions: After the integrity test, the filter cartridge and the housing are wet. To obtain storing conditions and prevent product dilution, the filter and its housing must be dried. The housing is therefore flushed with warm sterile air, which is evacuated on the upstream side as well as on the downstream side under sterile conditions. As water evaporates, warm air is cooled. The temperature is monitored on the incoming warm air and on the downstream side of the filter. When the temperature difference between the incoming air and the downstream side is smaller than a specific validated value, the filter and housing are dry. Drying process parameters are recorded in the data system for process consistency.

Closing Upstream and Downstream Valves: After storing conditions have been obtained, upstream valves (if defined in user requirement specifications) and downstream valve are closed (default position). This keeps the filter sterile. The housing (and its valves) is then disconnected from the test bench and may be used immediately or put on stock for later use.

Installing a Housing into a Process (Classical Stainless Steel): Because the valves ensure that the filter is sterile, only the connection must be sterilized. Sterilization can be conducted in two ways: either while steaming the whole process line (Figure 1) or by using steam through piping between a sterile housing and the already sterilized process line (Figure 2). The valves of the housing are then opened with compressed air piloted by the local PLC.

Large Housings: Large housings (e.g., multiround) might not be able to be displaced and put on the FMS test bench. So all required items (valves, sterile air, sterile water, and pressure sensors) are brought to the housing. Integrity testing is then started using a local touch panel.

Installing the Housing into a Single‑Use Bag Process Line: Single-use process lines commonly use single-use capsules that cannot be in-line steam sterilized and integrity tested using the FMS test bench. In such case, using a single-use steam-through connector (Figure 4) makes it possible to connect stainless-steel housings to any kind of single-use set-up.

Reducing Process Risk 
Many inspectors consider preuse poststerilization integrity testing (PUPSIT) a GMP requirement. However, some users and the PDA PUPSIT task force consider preuse poststerilization testing to increase process risk. The FMS test bench is an engineering way to reduce process risk commonly associated with PUPSIT. The FMS bench can be efficiently implemented only if a process line is designed from the start to integrate special designed housings. The economic viability of the FMS bench is based on the number of housings and number of tests performed per day.

More than just addressing the concerns of PUPSIT, the FMS method can significantly reduce downtime of a process line. No filter will ever block the production line because of integrity testing or required retesting preceded by time-consuming rewetting and error seeking. Based on experience with integrity testing failures and individual cost for process line downtime, a customer-specific return on investment can be calculated to determine whether the FMS is a cost-effective method.

Magnus Stering is product manager for Integrity Testing Solutions, Sartorius Stedim Biotech; magnus.stering@sartorius-stedim.com.

The post Preuse, Poststerilization Filter Integrity Testing for Single-Use and Stainless-Steel Installations appeared first on BioProcess International.

Increasingly efficient bioreactors allow biopharmaceutical manufacturers to achieve higher cell densities. That improved upstream efficiency has led to new purification challenges resulting from high product and contaminant concentrations as well as complex components. Therefore, harvest and clarification techniques are evolving to incorporate feed pretreatment, flocculation, and different filtration technologies such as normal-flow, tangential-flow, and depth filtration. The objective is to increase process capacities and filtrate quality, ultimately reducing biomanufacturing costs.

New strategies for clarification of recombinant proteins (in particular, monoclonal antibodies) and methods of pretreatment can improve clarification efficiency. Such approaches lead to better purity of the obtained filtrate, improving the overall efficiency of downstream purification steps that follow.

Traditional Methods
Using traditional expression systems, bioreactors can support cell densities ranging from 10 to 50 × 10⁶ cells/mL, which allows for product concentrations of ~2–3 g/L. To collect the active intra- or extracellular molecules at the end of such a cell culture, cells may be disrupted (intracellular products and most microbial systems) or discarded (extracellular products and most animal-cell systems). Generated cell debris typically fall below 10 µm in size.

A clarification filtration train clarifies those complex mixtures. Centrifugation or primary filtration is followed by secondary filtration (e.g., using tangential-flow and depth filters) before purification involving chromatography (Figure 1). With a solid load of 3–5% of the volume, such a filtration train results in a typical active product yield >85%.

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Figure 1: Clarification process for mammalian cell culture

Tangential-flow filtration (TFF) is used in microfiltration mode (open cut-off from 300 kD to 0.65 µm). It requires thorough process monitoring of pressures, feed flow, flux, and so on. Therefore, TFF is more delicate to use than normal-flow filtration (typically involving depth filters).

When centrifugation is used (often a stacked- disc centrifuge), an intermediate tank is necessary before secondary clarification because centrate flow will be discontinuous. Short-solid discharge phases (cells and their debris) break the centrate flow. Continuous-flow centrifugation can be applied, and downstream steps will have to accommodate its imposed flow rate.

Depth filters are traditionally made of cellulose fibers and an inorganic filter aid (diatomaceous earth) assembled in a lenticular or flat self- contained module (1). Such filters ensure a normal- flow separation of particulates and solutes through two main mechanisms: sieving and adsorption. Based on size exclusion, sieving retains rigid particulates at the surface and depth of a filter. Adsorption works independently from size. It is based on interactions between filtration media and particulates through ionic attraction and hydrophobic or electrostatic forces.For bioreactors with a volume <2,000 L, depth filters represent a significant cost advantage for primary clarification and remain the simplest option.

Today’s Challenges
A decade ago, 200-mg/L antibody concentrations in bioreactors were considered excellent. Today, bioprocessors aim for 10-g/L concentrations, with cell densities >20 × 10⁶ cells/mL. Higher densities necessitate more challenging purification steps (chromatography, TFF), particularly when solids are >10% of the process volume.

Increased biomass naturally leads to higher contaminant loads (2). During primary clarification, with a greater proportion of solids/particulates to eliminate, a filtration cake builds at the surface of traditional depth filters. Their capacity is greatly reduced (<50 L/m²) because their depth is underused for sieving and adsorption (Figure 2). The filter area needed for primary clarification thus becomes unsuitable for industrial use.

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Figure 2: The combination of inefficiently used depth media (a) and formation of additional resistance layer (c) reduces capacity. Optimization of media pore-size distribution using graded depth-filter media and particle-size distribution using pretreatment improves depth-filter performance with higher capacities and lower turbidity (b).

New methods are thus being developed for more efficient primary clarification, combining feed pretreatment with next-generation depth filters.

New Clarification Strategies
To improve the efficiency and capacity of primary clarification by filtration and include single-use technologies, two parameters can be modified and combined: particulate-size distribution in feeds and filtration media structure.

Feed pretreatment is a well-known technique widely applied to wastewater treatment to build a “sludge” and remove biomass. In bioprocessing, pretreatment of bioreactor output causes a shift in average particulate size (>10 µm), which eases particle removal by normal-flow filtration on new depth filter media.

Determining the right pretreatment agent depends on the nature of the protein of interest, contaminants in the feed, and further purification steps downstream. To assess pretreatment efficiency during development, several parameters can be measured: e.g., turbidity in supernatant and particulate-size distribution (for which a Malvern Mastersizer analyzer is useful). Filter capacity is also assessed for both clarifying filters and downstream filtration (e.g., in secondary clarification or bioburden reduction). Product concentration and yield are also indicators for assessing the effects of a pretreatment step on product quality.

Pretreatment options include acids such as acetic acid; salts such as (NH₄)₂SO₄, K₂SO₄, and KH₂PO₄; cationic polymers such as chitosan; and other polymers such as pDADMAC and polyethylene glycol (PEG) (3, 4).

Acid treatment is the simplest method. The pH level of a process solution is lowered to 5 (optimized during trials) after addition of an acid, thus modifying the charges of the solutes and leading to aggregation or precipitation of medium-sized particulates (20–30 µm), which are then retained by filtration media. From a regulatory perspective, no agent removal is necessary because only pH is modified, and it is generally readjusted after filtration.

Using salts for protein purification by precipitation is a broadly applied method in biochemistry. The salts interact at the surface of contaminants, decreasing their solubility in the solvent (water) and leading to their precipitation. However, both this method and the pH adjustment described above can denature a protein of interest, thus decreasing its stability and/or yield.

Cationic polymers lead to flocculation. A flocculation agent (the polymer) acts as a “binder” to aggregate contaminants present in a solution, causing formation of a floc (Figure 3). Polymers are nonspecific, binding cell debris, host-cell proteins, nucleic acids, and other contaminants. Flocs are generally larger (30–60 µm) than particulates obtained through acidification. The success of a flocculation step depends mainly on polymer dosage. Removal of that polymer has to be validated because, from a regulatory perspective, free polymer is considered a contaminant.

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Figure 3: Floc formed in a bioreactor and mechanism of flocculation

Depth Filters: Filters for primary clarification of pretreated feeds are constructed to optimize use of their depth (sieving) while keeping a good adsorption capacity. Clarisolve depth filters from EMD Millipore are specifically designed for that purpose (5). The filter media are made of polypropylene with cellulose fibers and/or diatomaceous earth, depending on the grade.

Improved capacity of Clarisolve filters eliminates the need for primary clarification by centrifugation, which reduces capital investment and equipment maintenance while allowing for implementation of single-use technologies. In addition, primary and secondary clarification may be condensed into one step depending on pretreatment efficiency. That can significantly reduce both footprint and depth filtration area — by three to five times — from those of traditional media (Figure 4).

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Figure 4: Comparing filtration area required for primary clarification of treated (acid precipitation) and untreated feed

Required flushing volumes before processing also may be significantly reduced, depending on the nature of the media. For example, the layers of polypropylene media that make up Clarisolve 60HX-grade filters reduce flushing volumes by ≤90% (down to <25 L/m² to reach TOC ≤ 3 ppm) compared with media made of cellulose fibers and inorganic filter aids. This can contribute to a more efficient and economic clarification of high–cell- density bioreactors using largely single-use technologies and improving the efficiency of downstream purification steps.

Breaking the Bottleneck
Increasingly efficient bioreactors are enabling higher cell densities. Treatment of their output requires new clarification strategies to increase the efficiency of downstream purification. New clarification methods can reduce filtration area, time, and steps necessary for filter preparation (including flushing). And cleaning requirements can be minimized with single-use technologies. Specific strategies include feed pretreatment and using a macroscopic floc and filtering on media developed for optimum depth use. Compared with a traditional filtration train, these new strategies can improve filtrate purity and increase process capacity to ameliorate the downstream bottleneck.

References
1 Wang A, Lewus R, Rathore AS. Comparison of Different Options for Harvest of a Therapeutic Protein Product from High Cell Density Yeast Fermentation Broth. Biotechnol. Bioeng. 94(1) 2006: 91–104.

2 Zhao X, et al. Developing Recovery Clarification Processes for Mammalian Cell Culture with High Density and High Solid Content. 243rd ACS National Meeting and Exposition, San Diego, CA, 2012.

3 Riske F, et al. The Use of Chitosan As a Flocculant in Mammalian Cell Culture Dramatically Improves Clarification Throughput Without Adversely Impacting Monoclonal Antibody Recovery. J. Biotechnol. 128(4) 2007: 813–823.

4 Kang YK, et al. Development of a Novel and Efficient Cell Culture Flocculation Process Using a Stimulus Responsive Polymer to Streamline Antibody Purification Processes. Biotechnol. Bioeng. 110(11) 2013: 2928–2937.

5 Singh N, et al. Clarification of Recombinant Proteins from High Cell Density Mammalian Cell Culture Systems Using New Improved Depth Filters. Biotechnol. Bioeng. 110(7) 2013: 1964–1972.

Sarah Le Merdy is a field marketing specialist in core field marketing for purification and BMT Process Solutions at Merck Millipore, sarah.lemerdy@merckgroup.com. Clarisolve is a registered trademark.

The post Evolving Clarification Strategies to Meet New Challenges appeared first on BioProcess International.

To date, the majority of recombinant monoclonal antibodies (MAbs) have been produced by mammalian cells. During such production processes, the potential risk of entrained viruses must be critically considered (1). Contamination can arise from animal cell lines or from adventitious viruses introduced during manufacturing. To ensure the viral safety of biotechnology products, companies can take four complementary approaches (2, 3):

• Using animal-component–free raw materials wherever possible
• Virus testing of master cell banks
• Virus testing of unprocessed harvest
• Performing downscale virus validation studies on selected process steps to demonstrate the capacity of the process to remove and inactivate viruses.Image may be NSFW.
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In most antibody-purification processes, the first viral clearance step is a low-pH inactivation (pH <4) conducted after protein A affinity capture chromatography. Precipitation of impurities often occurs in this low- pH treatment and subsequent conditioning, and filtration is required afterward before further processing in chromatographic polishing steps. Depth filters are commonly used (Figure 1).

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Figure 1: Antibody purification process with adsorptive depth filtration as a viral clearance step orthogonal to low-pH inactivation and virus filtration; the additional virus removal step in this process presumably enables replacement of anion-exchange chromatography as a polishing step while maintaining the virus safety of the process. Thus, a two-column purification process could be implemented (4, 9).

Many processors apply charged depth filters to make better use of that filtration step. Charged depth filtration is a scalable, well- characterized unit operation commonly used in blood-plasma fractionation processes, providing viral log clearance of ≤6 LRV (log₁₀ reduction value) (7, 8).

Previous investigations of antibody processes have demonstrated the ability of positively charged depth filters to reduce host-cell proteins (HCPs) and DNA (4) and to effectively remove viruses (1, 5, 6). Some investigators postulate that the virus retention can be attributed to anionic adsorptive effects of mammalian viruses on positively charged filter media (5). Such recommended model viruses for downscale virus validation studies as xenotropic murine leukemia virus (X-MuLV) and minute virus of mice (MVM) have a negative net charge at neutral or basic pH. Consequently, they bind to the positively charged surface of depth filters.

Our objective was to demonstrate the capabilities of three different adsorptive filters to remove viral contamination under typical process conditions. We tested laboratory-scale modules of these 3M filter types: Zeta Plus EXT 60ZA05A, Zeta Plus EXT 90ZA05A, and the functionalized nonwoven (FNW) Emphaze anion- exchange (AEX) Hybrid Purifier. The chromatographic characteristic of the latter filter comes mainly from its comparably high content of positively charged quaternary groups captured within a non-woven matrix. For these downscale virus retention studies, we used the mammalian viruses X-MuLV and MVM.

Materials and Methods
For these adsorptive depth filtration studies, we used ÄKTA Explorer 10 systems with Unicorn software from GE Healthcare. All buffers were 0.2-µm filtered before use. Table 1 lists our adsorptive test filters with anionic exchange functionality.

Load Material and Virus Assay: Rentschler scientists performed virus- spiking experiments at Charles River Biologics Testing Solutions in Cologne, Germany, using X-MuLV and MVM as model viruses (Table 2). Challenge materials were prepared from processed MAb-containing harvest.

Table 1: Overview of used filter types and process parameters used

Table 2: Virus models used for the virus clearance study

Culture harvest was clarified using a 3M Zeta Plus encapsulated 16EZA system with E16E01A60SP02A capsules (3 × 0.23 m²), 0.2-µm filtered and purified using protein A affinity chromatography. Column elution was achieved using a phosphate–citrate– Tris buffer system, and the eluate was adjusted to pH 3.5 and diluted to a MAb concentration of 5 g/L. Before the adsorptive depth filtration runs, the protein solution was divided and each sample load adjusted to desired loading conditions (pH value and sodium chloride concentration) (Table 3). Those adjustments were made using a phosphate–citrate–1 M Tris buffer and 4M NaCl solution, leading to a final MAb concentration of 3.75 g/L.

Table 3:  Experiments carried out with all three depth filters

To exclude influence of test items on cell growth or virus replication, Charles River Biologics Testing Solutions also performed cytotoxicity and viral interference assays. The same test facility also provided virus stock solutions and further analyzed samples using 50% tissue-culture infective dose (TCID₅₀) assays. The detection limit of such assays depends on the volume of sample that is incubated with indicator cells. Those cells were cultivated for a specific incubation period and inspected microscopically for virus-induced changes in their morphology.

Experimental Methods: Viral clearance experiments were performed with the three adsorptive filter types using three different buffer conditions, each in duplicate (Table 3). Virus- reduction factors of X-MuLV and MVM were determined for these conditions

• phosphate–citrate–Tris buffer at pH 5.5 with 0 M NaCl (“pH 5.5 low salt”)
• phosphate–citrate–Tris buffer at pH 7.0 with 0 M NaCl (“pH 7.0 low salt”).

Those buffers represent appropriate conditions for further processing in a MAb purification. Additionally, viral reduction of X-MuLV (80–100 nm) was determined in a phosphate–citrate–Tris buffer at pH 7.0 with 1M NaCl (“pH 7.0 high salt”) to minimize ionic interactions between viruses and the adsorptive filter material. By subtracting the high-salt condition’s virus LRV from that of the low-salt condition, we calculated the viral clearance related to adsorption. Because MVM (20–25 nm) is much smaller than the nominal pore size of these filter media (0.2–0.8 µm for the Zeta Plus VR series), no additional filtrations were run under high-salt conditions. We excluded the mechanical effects for retention of MVM.

Duplicate filtrations were run in parallel with two chromatography systems, so 180 mL of the affinity- purified and conditioned MAb solutions were spiked with 5% (v/v) (9 mL) of nonconcentrated, ultracentrifuged, and prefiltered virus stock solution (0.45-µm filter for X-MuLV and 0.1-µm filter for MVM). After mixing, a sample was withdrawn from the spiked pool and immediately analyzed for the virus titer (“load sample”). For X-MuLV, a second load sample was stored until the end of the filtration (“hold sample”). MVM is known to be physicochemically resistant (1).

Analysts loaded 220 L/m² filter area (84 mL) of the starting material at 158 L/m²/h (1 mL/min) to the equilibrated adsorptive depth filters. Subsequently, they flushed those filters with 63 L/m² (24 mL) equilibration buffer. At the end, they analyzed the total filtrate virus titer (both flow-through and flush, referred to herein as the “filtrate sample”) and determined the titer of the drawn “hold sample.” They calculated virus log-reduction values as described in Equation 1:

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in which C_L = virus concentration of the load, V_L = volume of the load, C_F = virus concentration in the filtrate, and V_F = volume of the filtrate.

Results and Discussion
From cytotoxicity and viral- interference assays performed by Charles River Biologics Testing Solutions (data not shown), we obtained confirmation that no test items influenced cell growth or virus replication. The analysts also determined suitable sample dilutions for performing the TCID₅₀ assays.

Removal of X-MuLV: Under low-salt conditions at pH 5.5 and pH 7.0, we observed no positive indicator cells of X-MuLV (TCID₅₀ assay) in filtrate fractions from all three tested filters. Excellent virus removal was obtained, at least in the range of 5.0–5.8 LRV (Table 4). By contrast, under high-salt conditions at pH 7.0, we determined virus removal of 1.2 LRV (Emphaze AEX FNW) and 2.7 LRV (Zeta Plus 60ZA05A and 90ZA05A) filters. Because high-salt concentrations suppress electrostatic interactions, we used such conditions to investigate virus removal without attractive electrostatic interactions. The low value for the Emphaze AEX FNW filter shows no significant virus removal capacity at high-salt condition; the assay’s standard deviation is ±1 LRV.

Table 4: X-MuLV clearance results for the three charged depth filters tested

The LRV difference between high- and low-salt conditions can be traced back to virus adsorption on positive- charged filter media. To understand this adsorptive X-MuLV retention, we calculated a revised LRV with values obtained under high-salt conditions (pH 7.0) subtracted from the value under low-salt conditions (pH 7.0 and pH 5.5) (Table 5, Figure 2). Doing so provided values of 2.2–3.1 LRV for the Zeta Plus depth filters. The Emphaze AEX FNW filter showed revised LRVs that were still high at 4.7 and 4.0 (Table 5, Figure 2). So the main retention mechanism of X-MuLV on the latter at low ionic strength can be attributed to ionic adsorption.

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Figure 2: Revised log-reduction values for X-MuLV indicate virus clearance is based on anionic retention mechanisms.

Table 5: Revised log-reduction values for X-MuLV

Revised LRVs were lower for Zeta Plus 60ZA05A and 90ZA05A because of the persistent retention (2.7 LRV) at high-salt concentration. That outcome is probably related to hydrophobic interactions and/or marginal mechanical retention. Previous viral clearance studies using Zeta Plus 90ZA filters had already shown that hydrophobic interactions are involved in virus retention at high ionic strengths (6). That phenomenon can be explained by the net charge of X-MuLV, which is nearly zero at pH 7.0 because the heterogenic isoelectric point (pI) of its major capsid protein (p30) ranges from 6.1 to 6.6 (10). Additionally, the p30 protein has many hydrophobic amino acids, possibly presenting exposed hydrophobic moieties.

A minor retention by size may be attributed to convective entrapment that occurs (even if a filter’s pore size is larger than the virus) when viruses are transported to pores by convection but are unable to traverse them fully because of irregular internal structures (11). A further possible reason for LRVs of >1 in the presence of 1M NaCl is related to ionic strength, which is not sufficient to suppress entirely the positive zeta potential. Therefore, some positively charged spots inside the filter might lead to virus removal even at high NaCl concentrations.

With each virus load, two filtration runs were performed in duplicate to provide one value for the load’s total virus load and two values for the filtrates’ total virus load for each tested condition and filter. Reduction values (Equation 1), and a mean value were calculated. The measured total virus load of the filtrates is indicated with mathematical symbols for “equal or less than” (≤) where no virus positive cells were observed with the TCID₅₀ assay sample dilution.

Revised reduction values are calculated by subtracting the means of the reduction values under low-salt conditions (pH 5.5 and pH 7.0) from the means of the reduction values under high-salt conditions (pH 7.0). In the presence of 1M NaCl, electrostatic interactions are widely suppressed independently from the pH value. Therefore, the LRV from the low-salt conditions at pH 5.5 also can be subtracted from the LRV of the high-salt concentration at pH 7.0.

MVM Removal: We observed a very effective MVM removal (4.6–7.2 LRV) for all three depth filters at pH 5.5 and a clearance of 2.4–2.7 LRV at pH 7.0 (Table 6, Figure 3). The nominal pore sizes of all three filters are significantly larger (>0.2 µm) than the MVM virus (20–25 nm), so we believe that such virus-removal capacity is almost solely related to adsorptive retention (3, 12).

Table 6: MVM clearance results for the three depth filters tested

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Figure 3: Log-reduction values for MVM at pH 5.5 and pH 7.0

The pI of MVM is about pH 5 (13). So at pH 7.0, the virus should be negatively charged, and at pH 5.5 it should be nearly free of net charge. The higher MVM retention at pH 5.5 indicates that attractive electrostatic effects may play a minor role. It has already been shown that viruses are more effectively removed if operational pH values are near their pI (6).

To explain the mode of virus adsorption in more detail (proportion of ionic and hydrophobic interactions), additional studies can be performed with even higher salt concentrations and/or using organic compounds such as propylene glycol (PG) to suppress electrostatic and hydrophobic interactions. Such studies can improve understanding of the mechanisms responsible for virus retention. However, a detailed breakdown of adsorptive virus removal on a charged depth filter applied in an antibody-purification process that further includes pH virus inactivation and a nanofiltration (size exclusion) is unnecessary to evidence the orthogonality of those removal steps.

With each virus load, our analysts performed two filtration runs in double determination, leading to one value for the load’s total virus load and two values for the filtrates’ total virus load for each tested condition and filter. Reduction values (Equation 1) and mean values were calculated. Measured total virus load of the filtrates is indicated with mathematical symbols for “equal or less than” (≤) where no virus-positive cells were observed in the TCID₅₀ assay sample dilution.

Highly Efficient Virus Removal
We have demonstrated the capability of Zeta Plus 60ZA05A, Zeta Plus 90ZA05A, and Emphaze AEX Hybrid Purifier filters in a unit operation for viral clearance of a MAb purification process. A protein A– purified antibody solution was spiked with two routinely used mammalian model viruses (X-MuLV and MVM), and the downscale virus-retention study was performed under conventional process conditions.

For X-MuLV clearance by Zeta Plus 60ZA05A and 90ZA05A filters, values of 4.7–5.8 LRV could be obtained at pH 5.5 and pH 7.0. We attributed LRVs in the range of 2.2– 3.1 to electrostatic retention mechanisms. The residual virus clearance is probably attributable to hydrophobic or mechanical interactions. The Emphaze AEX FNW filter removed X-MuLV with LRVs of 5.1–5.8 at pH 5.5 and 7.0 (low ionic strength), so its high values of 4.0–4.7 could be attributed to electrostatic retention. The small, nonenveloped MVM virus was reduced efficiently by all tested filters at pH 5.5, with LRVs of 4.6–7.2. MVM clearance was less effective at pH 7.0 (LRV ~2.5).

We found that Zeta Plus and particularly the Emphaze AEX FNW adsorptive depth filters can be considered highly efficient virus- removal options under typical conditions in a MAb downstream process. Moreover, the excellent viral clearance potential of the Emphaze filter by adsorptive retention can be claimed as an orthogonal method for viral safety with the currently adopted virus-filtration step (size exclusion) and low-pH inactivation. The virus removal capability of these charged filters is complementary to other virus clearance steps in downscale validation studies of a MAb process. Just as in validating an anion-exchange chromatography step, the viral- reduction capacity of a charged depth filter must be determined under actual process conditions. So it has to be done individually for specific processes, with different amounts of process-related impurities (HCPs, DNA). We have shown this virus removal capability for our model antibody process using appropriate buffers and conditions.

The capability to cover the purpose of an anion-exchange polishing step with a positively charged depth filter will directly allow us to redesign our three–chromatography-step MAb purification platform into a more efficient and economical two-step process while maintaining product purity, quality, and safety.

Acknowledgments
The authors thank Pia Reimann, Peter Koklitis, and Carsten Erdmann at 3M for their project support and review of this manuscript.

References
1 Zhou JX, et al. Viral Clearance Using Disposable Systems in Monoclonal Antibody Commercial Downstream Processing. Biotechnol. Bioeng. 100(3) 2008: 488–496.

2 ICH Q5A (R1). Viral Safety Evaluation of Biotechnology Products Derived from Cell Lines of Human or Animal Origin. US Fed. Reg. 63(185) 1998: 51074.

3 Bergmann K, et al. Design and Performance of Viral Clearance Studies. BioProcess Int. 4(10) 2006: 56–62.

4 Faude A, et al. Virus Removal By Simple Depth Filtration in a Two-Step Downstream Process for MAb Production (poster). Recovery Conference, Rostock, 2014; http://rentschler.de/fileadmin/Downloads/ Rentschler_Poster_Recovery_201407.pdf.

5 Wang M. Zeta Plus™ VR Filters for Viral Reduction. BioProcess Int. 9(7) 2011: 62.

6 Zhou JX. Orthogonal Virus Clearance Applications in Monoclonal Antibody Production. Process Scale Purification of Antibodies. Gottschalk U, Ed. John Wiley & Sons: Hoboken, NJ, 2009; 169–186.

7 Revie D, et al. Novel Application Approaches to Obtain Maximum Viral Clearance from an Immunoglobulin Production Process. IBC’s Second International Symposium on Viral Clearance, 1998.

8 Application Brief. Cuno Zeta Plus®VR Filters for Viral Reduction in Blood Plasma Fractionation Processes. Cuno (3M): Meriden, CT, April 2002; http://multimedia.3m.com/ mws/media/418846O/zeta-plus-vr-filters-viral- reduction-blood-plasma-fractionation. pdf?fn=DOC00042%20-%20 LITCABZPVR1.E.pdf.

9 Müller D, et al. Taking MAb Purification to the Next Level. Europ. Biotechnol. Life Sci. Ind. 13, 2014: 76.

10 Mitra S, et al. Comparative Physicochemical and Biological Properties of Two Strains of Kilham Rat Virus, a Non-Defective Parvovirus. J. Gen. Virol. 61(Pt l) 1982: 43–54.

11 Trilisky, et al. Flow-Dependent Entrapment of Large Bioparticles in Porous Process Media. Biotechnol. Bioeng. 104(1) 2009: 127–133.

12 Hou K, et al. Capture of Latex Beads, Bacteria, Endotoxin, and Viruses By Charge- Modified Filters. Appl. Environ. Microbiol. 40(5) 1980: 892–896.

13 Ray S, et al. Chapter 26: Single-Use Virus Clearance Technologies in Biopharmaceutical Manufacturing: Case Studies. Single-Use Technology in Biopharmaceutical Manufacture. Eibl R, Ed. John Wiley & Sons: Hoboken, NJ, 2011: 311–322.

Mario Metzger and Anja Gerster are process engineers in technology development; Marcus Peiker is process manager, Sabine Faust is a process engineer, and Alexander Faude is associate director of downstream process development; Sybille Ebert is assicate director, and corresponding author Dr. Dethardt Müller (dethardt. mueller@rentschler.de) is vice president of technology development at Rentschler Biotechnologie GmbH in Laupheim, Germany. Dr. Sophie Winterfeld (swinterfeld@mmm.com) is senior engineer in application development for life sciences process technologies, and Nicole Mang is a senior sales account executive in life-science process technologies at 3M Deutschland GmbH in Neuss, Germany.

The post Evaluating Adsorptive Filtration As a Unit Operation for Virus Removal appeared first on BioProcess International.

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EMD MILLIPORE (WWW.EMDMILLIPORE.COM)

In the manufacture of monoclonal antibodies (MAbs), the first purification step following harvest clarification is normally protein A affinity chromatography because of its high selectivity for IgG and high process yield (1, 2). At this stage, a MAb is eluted from a protein A ligand at low pH and then held or adjusted to a low pH (pH ≤ 3.8) for a given amount of time before pH adjustment, usually ≥30 minutes, in a virus inactivation (VI) step targeted at retroviruses (3, 4). After VI, product pools are adjusted to a pH that is appropriate for further downstream unit operations, which may include one or several additional modes of chromatography, virus filtration, and final formulation (5).

During the VI and subsequent pH adjustment steps, operators commonly observe a significant amount of precipitation and turbidity in their product pools (2). Precipitants require larger sterilizing-grade filter areas after VI. Components that are generally responsible for precipitation at low pH (or during pH adjustment) include product-related species such as high– molecular-weight aggregates and process-related molecular impurities such as host-cell proteins (HCPs) and DNA (3, 6). Increased process-related impurities such as HCPs and DNA can be related to cell type (e.g., bacterial/yeast lysate will produce more impurities than will Chinese hamster ovary (CHO) cells) (7). It also can come from increased cellular debris that can be attributed to excessively low cell viability at the time of harvest (8).

Reported solutions to the precipitation problem include increasing the surface area of the sterilizing-grade filter and using inline prefiltration (6). However, because of potentially slow kinetics, the end point of a precipitation event may be difficult to characterize, thus leading to unpredictability of filter-area requirements and even possible filter failure (6).

Another option is using adsorptive depth filtration (ADF) containing diatomaceous earth (DE) to remove precipitants in pH-adjusted protein A eluates. Kandula et al. cautioned that such DE use in downstream processing following protein A may be viewed as unfavorable because of the potential for leachables introduction to process streams (6). However, that concern is minimal for processes in which a bind– elute chromatography or ultrafiltration/ diafiltration (UF/DF) step is used downstream of ADF. Those steps should be able to effectively clear small leachate species (that are not already removed by ADF flushes) by the bind–elute flow-through action or UF/DF permeation (9).

The capacity of depth filters to remove turbidity and protect sterilizing filters has been demonstrated, as have the ability of depth filters to reduce soluble process-related impurities. Yigzaw et al. thoroughly investigated the ability of depth filters to reduce the amount of HCP in a MAb process through optimization of the harvest clarification step (10). For that study, turbidity (measured by UV absorbance at 410 nm) was used to monitor the effectiveness of ADF in reducing HCP. The work showed an ability of depth filtration to reduce HCP in cell-culture harvest fluid. In terms of molecular impurity removal downstream of the protein A capture step, Haverstock et al. characterized varying levels of HCP removal from pH-adjusted pools as a function of pH (11).

We initially investigated ADF to supplement removal of HCPs in a MAb purification process, adding a depth filter to the existing process along with the normal sterilizing-grade filter following VI. Additional testing of the depth filter helped us characterize the reduction of other process-related impurities, including residual CHO DNA and residual protein A ligand. We performed these tests at both laboratory (10 L) and production (1,000 L) scales. Upon implementing ADF into this process, we also investigated the feasibility of removing a subsequent column step.

Materials and Methods
Monoclonal antibodies A (pI 8.15– 8.65) and B (pI 8.6–8.9) are IgG1 MAbs produced by CHO cells.

Cell Culture: CHO cells producing either MAb-A or MAb-B were cultured, expanded, and inoculated into 10-L to 2,000-L bioreactors with proprietary media free of animal- derived components. We clarified the 10-L cultures using ADF and sterile filtration and supernatant from 130-L, 1,000-L, and 2,000-L reactors using centrifugation, ADF, and sterile filtration. All reactors were harvested on day 15 after inoculation and/or within 24 hours of cell viability falling below 70%.

Purification: Following clarification, MAb-A and MAb-B cell culture supernatants were loaded onto a MabSelect SuRe protein A affinity chromatography column from GE Healthcare. We eluted both MAbs at low pH. Then we adjusted the pH of MAb-A eluates using 2 M acetic acid to 3.7 and held them for an hour for VI of enveloped viruses before adjusting pH to 6.5 (unless otherwise specified) using 2 M tromethamine (Tris). We adjusted the pH of MAb-B eluates to 5.0 using 2 M Tris immediately upon elution from protein A, then stored them at 2–8 °C and later adjusted pH to 6.5, 7.0, or 7.5.

Adsorptive Depth Filtration: Following pH adjustment, the MAbs were loaded onto either a Millistak+ D0HC ADF prefilter or a Millstak+ X0HC ADF terminal filter (both from EMD Millipore) at a flux of 100 L/m²/h (LMH) (unless otherwise specified) to a target loading of 100 L/m² (unless otherwise specified) based on the area of the X0HC filter. We used device areas of 23 cm² for small-scale trials and 270 cm^2, 540 cm², or 1.1 m² device areas for intermediate- and process-scale confirmations. During depth filtration, differential pressures were monitored as a function of loading, and we calculated filter resistance as the ratio of differential pressure to flux. Figure 1 shows our experimental set-up for these ADF experiments.

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Figure 1: Diagram of experimental set-up used to assess adsorptive depth filtration (ADF) in MAb-A and MAb-B processes following protein A; in this diagram “ADF” refers to either the terminal ADF, ADF prefilter, or a filter train consisting of an ADF prefilter and ADF in series. Unless otherwise specified, flux onto an ADF is standardized to 100 L/m²/hour.

Anion-Exchange Resins and Membranes: Anion-exchange (AEX) resin 1 consisted of quaternary amine ligand on a highly cross-linked agarose base matrix. AEX resin 2 consisted of a quaternized polyethyleneimine ligand on a cross- linked poly(styrene-divinylbenzene) base matrix. AEX membrane 1 is a stabilized reinforced cellulose membrane with a quaternary ammonium ligand. AEX membrane 2 is an ultrahigh–molecular-weight polyethylene membrane with a primary amine ligand.

Host-Cell Protein: Unless otherwise specified, we determined HCP concentrations by immunoassay with a third-generation CHO HCP enzyme- linked immunosorbent assay (ELISA) kit from Cygnus Technologies, carrying out the ELISA method according to the manufacturer’s protocol. For Experiment 3, we used a sandwich ELISA to measure CHO HCP. Capture antisera bound the HCP, which was then detected using biotinylated antisera and extravidin–horseradish peroxidase. This method used ABTS as a chromogenic substrate. Samples of unknown HCP concentration were interpolated from a standard curve.

Residual Protein A Ligand: We measured residual protein A concentration by ELISA as well, with a MabSelect SuRe kit from Repligen. Briefly, a capture antibody bound MabSelect SuRe protein A, which is then detected using biotinylated rabbit antibodies against protein A and streptavidin–horseradish peroxidase. We used tetramethyl benzidine (TMB) as the chromogenic substrate. Samples of unknown concentration were interpolated from a MabSelect SuRe protein A standard curve.

Residual CHO DNA: For Experiment 3, our procedure for determining residual CHO DNA concentration was based on quantitative polymerase chain reaction (qPCR) and an ABI PRISM 7900HT sequence detection system. The CHO qPCR assay was designed to detect a CHO retroviral sequence that may be present in CHO cell-line products. The amount of CHO DNA present in each sample was interpolated from another standard curve.

For Experiment 6, our procedure for determining residual CHO DNA concentration was based on the PrepSeq DNA sample preparation kit and resSEQ quantitative CHO DNA kit from Life Technologies. For real- time PCR analyses, we use an oligonucleotide probe containing both a fluorescent reporter dye and a quencher. When the probe is intact, the fluorescence is quenched. If a target sequence is present, the probe anneals and the reporter dye is subsequently cleaved from the probe by the 5′-nuclease activity of Taq DNA polymerase as the associated primer is extended, generating a fluorescence signal. In each run, we generated a standard curve from known amounts of CHO DNA and calculated the amount of residual CHO DNA in our samples from that standard curve.

Results and Discussion
Experiment 1 — Initial Characterization of ADF for MAb-A:

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Table 1: IgG recovery and host-cell protein (HCP) level at an adsorptive depth filter (ADF) volumetric loading of 160 L/m² as a function of supernatant pH

We loaded protein A eluate of MAb-A (after adjusting to pH 3.5, 6.5, or 7.5) onto a terminal ADF to ≥160 L/m² and took in-process samples across these filtrations at 20-L/m² intervals. At all loading levels, process streams adjusted to pH 6.5 and 7.5 contained lower HCP levels than the pH 3.5 stream (Figure 2), which is probably the result of more negatively charged impurities adsorbing to the ADF filter media. IgG recovery levels increased with decreasing feed pH, which is also consistent with the existence of an interaction with the positive charge contained in the ADF filter (Table 1).

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Figure 2: Host-cell protein (HCP) content in adsorptive depth filtrate samples remains lower at higher supernatant pH, which is consistent with an interaction between negatively charged HCP and positively charged ADF. Control uses only 0.2-µm membrane filtration.

Experiment 2 — Scale-Up Confirmation of MAb-A ADF: We performed a series of three ADF experiments using intermediate-scale devices. The first such experiment used a 270-cm² terminal filter alone; however, due to increased turbidity of the feed material (because of increased IgG and impurities concentration in the protein A eluate), the ADF reached an excessively high differential pressure (>20 psi) and could not be loaded beyond 66 L/m² (compared with 160 L/m² previously). We took in-line samples at 20-L/m² intervals during this experiment.

In the second experiment, we mitigated the pressure increase by using a looser-grade prefilter, which enabled removal of larger particles and reduced plugging on the denser terminal ADF filter. During this trial, we loaded both the prefilter and terminal ADF filter to 93 L/m² and took in-process samples at 20-L/m² intervals. We observed a modest pressure increase with this filter combination, but it was well within processing tolerances. In our third experiment, we loaded starting material across a 23-cm² ADF prefilter device alone (at 238 L/m²) to demonstrate the capacity of the ADF prefilter.

Figure 3 illustrates resistance as a  function of loading for all three trials using these ADF filter trains. We measured HCP across all three filtrations, and Figure 4 shows the results. Starting HCP levels and conductivity (16 mS/cm, 6 mS/cm) in the ADF load material were higher in the scale-up run than in the previous small-scale experiments, so the sample fraction HCP levels also were higher. However, we confirmed HCP content in the depth-filtered fractions to be lower than that measured using dead-end (0.2-µm) filters alone (Figure 4). We observed little difference in HCP as a function of loading with the addition of the ADF prefilter. Additional analyses showed that very little HCP was removed across the ADF prefilter alone, as expected, because of the lower charge density of that protective filter. We did observe a difference in HCP content between our starting loading material and the 0.2- µm only control, which may be related to assay variability or removal of precipitation caused by adjustment to pH 6.5.

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Figure 3: Resistance as a function of loading for terminal adsorptive- depth filtration (ADF), ADF prefilter, and ADF filter train shows that terminal ADF alone reaches higher pressure than when a prefilter is added in-line

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Figure 4: Plot of host-cell protein (HCP) content in adsorptive depth filtrate samples shows that the terminal ADF reduced HCP content relative to that of starting load material, non-ADF (0.2-µm filter only) control, and ADF prefilter alone.

Experiment 3 — Process-Scale Verification of ADF for MAb-A: Here, we implemented the ADF process demonstrated at intermediate scale above for pilot-scale manufacturing in a 130-L bioreactor. For small-scale verification of the planned larger-scale depth filtration, we filtered a sample of protein A eluate across the ADF prefilter (23 cm²), then pooled the results and filtered them across a terminal ADF (23 cm²) to a loading of 223 and 233 L/m² for each filter, respectively.

We depth filtered the remaining bulk of the eluate pool using a 270-cm² ADF prefilter, pooled the results, and then filtered them using a 540-cm² terminal ADF filter. Filters operated at fluxes of 186 and 93 LMH and to loadings of 238 and 119 L/m² (for the prefilter and terminal filter, respectively) without observable pressure increases across either filter. We took samples at intervals throughout this process and found that all in-line and pooled ADF samples exhibited HCP levels below the level of detection (LOD) of our assay. The ADF prefilter and terminal ADF filter reduced the CHO DNA levels from 34.6 pg/mg to 0.8 pg/mg. Residual protein A was reduced from 1.14 ng/mg to a level below the LOD (0.4 ng/mg) of the assay.

We found DNA, residual protein A, and CHO HCP in the ADF pool to be very similar to that of the pool from the subsequent AEX flow-through (FT) step: 0.8 pg/mg, <0.4 ng/mg, and <38 ng/mg for ADF; <0.8 pg/mg, <0.4 ng/mg, and <28 ng/mg, respectively for AEX FT. Based on this observation, we designed and executed follow-up experiments to assess whether the AEX column would be necessary for removal of these impurities in a MAb-A purification process that included ADF after protein A capture.

Experiment 4 — MAb-A Process Comparison With and Without Depth Filtration: We designed another set of experiments to determine the contribution of the ADF train in removal of impurities in MAb purification. First we split the adjusted protein A eluate into two separate process streams, one including ADF and the other without ADF. We loaded the stream with ADF (including a prefilter at a 1:1 ratio) to 126 L/m² and filtered the stream without ADF with a 0.2-µm filter alone. Both streams were processed separately through subsequent downstream AEX FT and bind–elute (BE) polishing chromatography steps. We sampled pools from all process intermediates.

HCP levels were lower throughout the process in the stream containing ADF and were 0.87 log10 lower following the final polishing step than for the process without ADF (Table 2). We estimated the level of MAb-A recovery across the X0HC depth filter in this experiment to be 94%.

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Table 2: Host-cell protein (HCP) profile through a MAb-A purification process with and without adsorptive depth filtration (ADF)

Experiment 5 — MAb-A Process Comparison With and Without an AEX Column: We loaded adjusted protein A eluate material across a terminal ADF filter to 98 L/m², this time splitting the ADF pool into two streams to compare processes with and without an AEX FT step. We took pool samples after the remaining steps of the purification process for both streams and measured MAb-A recovery across the depth filtration to be 83.6%. We attributed that low recovery level to an early termination of the recovery flush through the ADF filter.

Table 3 shows that the ADF step removed 2.6 log₁₀ of HCP, whereas the subsequent AEX FT step had negligible impact on HCP removal. The final HCP values after BE polishing chromatography were almost identical across the streams both with and without AEX FT. We observed a similar trend for residual protein A when tracking that throughout the process. These results suggest that using an ADF operation after protein A capture rendered the use of a subsequent AEX chromatography step redundant for clearing residual CHO HCP and residual protein A ligand impurities.

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Table 3: Comparing residual Chinese hamster ovary (CHO) host-cell protein (HCP) and protein A levels for processes that either include or omit anion-exchange flow-through chromatography, both having an adsorptive depth filter (ADF) following pH adjustment, virus inactivation (VI), and pH adjustment

Experiment 6 — MAb-A 1,000-L Scale-Up Process Verification: From a 1,000-L scale run, we filtered an adjusted protein A eluate using a 0.55-m² ADF prefilter (to 112.4 L/m_²) and a 1.1-m² ADF filter (to 56.2 L/m²). To verify the results obtained in Experiment 5 at a larger scale for HCP, residual CHO DNA, and residual protein A ligand removal in a MAb-A process, we tested material from a 1,000-L purification process according to Figure 5. Table 4 shows that all levels were reduced across the ADF and that including an AEX step had a negligible effect on those impurity levels.

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Figure 5: During a 1,000-L bioreactor purification run, material was taken for small- scale experiments to demonstrate the effect of ADF on impurities removal in the MAb-A process both with and without AEX.

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Table 4: Residual CHO DNA, protein A, and HCP assay results from 1,000-L bioreactor processing

Experiment 7 — Comparison of ADF with AEX Resins and Membranes in MAb-B Process: For a second antibody (MAb-B), we compared a purification process that included a terminal ADF step with processes including either AEX chromatography or an AEX membrane filtration following the capture step. We purified protein A eluates adjusted to pH 6.5, 7.0, or 7.5 over one of the following matrices: ADF, AEX Resin 1, AEX Resin 2, AEX Membrane 1, or AEX Membrane 2. Table 5 lists the resulting ADF/AEX matrix volumes, loading/flush flow rates, loading levels, and recoveries. Figure 6 shows assay results from in-line samples taken during the load and recovery flush. Pool recovery is based on an IgG mass balance.

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Table 5: Comparing run parameters for adsorptive depth filter (ADF) and anion-exchange (AEX) matrices

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Figure 6: ADF showed higher clearance of residual CHO HCP than two AEX resins and membrane adsorbers; because scale could not be normalized to match other datasets, data from AEX membrane 2 is not shown here (at pH 7.5 with 634 mg MAb-B/0.08-mL membrane, CHO HCP was measured at 66 ng/mg).

Those results show that terminal ADF provides greater HCP reduction than the other matrices tested. Recovery levels across the ADF filter at pH 7.5 were similar to those seen for both AEX resins and slightly lower than those measured for both AEX membranes. Such levels of clearance (about one additional log of HCP ng/mg clearance for most comparisons) are similar to MAb-A clearance levels in Table 3.

Discussion
The levels of process-related impurity removal we measured for residual CHO HCP, residual CHO DNA, and residual protein A ligand in samples following ADF were at least as low as (or lower than) those seen in AEX flow-through samples for two MAbs. These results suggest that adding a depth filter postcapture to a MAb purification process could make the AEX step redundant for removal of those impurities.

Observed recovery levels across the depth filter were slightly lower than those seen across the AEX resins used in both MAb-A and MAb-B processes. This is expected: In addition to the common charge interactions for both technologies, depth filtration can bind IgG or process-related impurities through hydrophobic interactions, which could be responsible for additional IgG adsorption (10).

Because of developmental time constraints and our objective to limit the extent of change from the original process, we did not optimize pH of the protein A eluate or pH and ionic strength of the buffer used for equilibration and recovery flushing in depth filtration for the terminal ADF step of MAb-A. Additional improvements in recovery and/or HCP reduction might be realized if those parameters are characterized more fully. We considered only 100 L/m² loading levels on the terminal ADF filter at a 100-LMH flux to maximize impurity removal. Additional gains in efficiency and productivity could be realized at higher levels of loading and flux, as shown by Haverstock et al., who found little change in HCP across higher flux levels (11).

Virus Safety Considerations: In addition to molecular impurity clearance of molecular species such as CHO HCP and DNA, AEX FT often is used to support virus safety in MAb processes. Our work demonstrates that ADF is suitable for removal of CHO HCP, CHO DNA, and residual protein A ligand; however, fully replacing AEX with ADF presents many challenges for viral clearance. With all other technologies that are claimed as virus-removal strategies (as opposed to virus inactivation), associated integrity tests must indicate the ability of a device to perform the virus removal function. For example, membrane filters have a corresponding postuse integrity test. And for chromatography columns, associated preuse specifications for asymmetry and height equivalent to a theoretical plate (HETP or plate height) indicate that flow distribution is uniform through the media and no resin-bed cracking is present to cause bypass of the fluid stream through those media.

If ADF were to be claimed in a viral-safety strategy, it would need a similar approach to ensuring virus- removal functionality. Current ADF manufacturing technology ensures that the filters can remove large particulates from relatively impure streams, where the effects of variability among device lots is minimal. Manufacturing technologies for virus-removal media are designed to much higher standards, however, and include rigorous lot- release testing specific to viral clearance. By contrast, no virus- removal claims are made for ADF. Providers of ADF media would need a testing strategy to ensure consistent virus clearance. The potential utility of ADF to remove viruses in a MAb process has been demonstrated (12, 13), but because of these challenges, the responsibility would be on end users to ensure that acceptable strategies are in place to test, validate, and confirm integrity within their own processes.

In a recent study, Iskra et al. (14) demonstrated the ability of ADF to lower HCP levels in a protein A pool and enhance removal of xenotropic murine leukemia virus (xMuLV) across a downstream AEX chromatography step. Their work demonstrates that even if ADF cannot be validated for virus removal, it could increase robustness of the downstream virus-removal capabilities of an AEX step.

Scalability: We demonstrated scalability of ADF filters from bench to pilot scale (1,000 L) during our initial phases of testing, with similar levels of recovery and CHO-HCP removal across the depth filtration steps at each scale. No immediate scale-up issues were observed, but future verification will be necessary. For example, recovery flush volume is one parameter that could affect recovery levels and thus will need to be optimized for future larger-scale runs.

Different Molecules: We have demonstrated the utility of ADF after VI for two MAb molecules to date. Our initial data suggest that an ADF process could be used as a replacement for an AEX FT step for removing HCP, residual protein A, and residual CHO DNA in MAb processes. Future studies will characterize and investigate not only additional molecules, but also the critical parameters for the ADF unit operation. Once those parameters are identified, a more in-depth characterization of the design space will help establish and quantify the levels of risk associated with an ADF step.

Leachables/Extractables: We have also considered the risk of introducing leachables into product streams. Leachables and extractables that have been characterized for depth filtration include organic carbon, metals, inorganic salts, and fibers (9). Upon discussion and risk assessment, we concluded that transmission of those leachables through the downstream process into final drug products was unlikely because

• the recommended initial flush volume for the depth filter specified in our purification process is predicted to remove a majority of leachables (15)
• both MAb-A and MAb-B ADF steps are followed by both BE chromatography and UF/DF steps.

The low-pH hold occurs early in a typical MAb purification process, which could allow most manufacturers the option of adding a depth filter. But the risk of leachables transmission through the process probably will be molecule and/or platform specific.

Promising Results, More Work to Do
We demonstrated the usefulness of terminal ADF to remove HCP, residual CHO DNA, and residual protein A ligands after the capture step in a MAb process for two molecules at multiple process scales. Our data suggest that adding terminal ADF to a MAb purification process could eliminate the need for AEX FT in that process. However, further characterization and verification of the ADF step will be necessary, and some outstanding questions will need to be addressed in future experiments.

References
1 Low D, O’Leary R, Pujar NS. Future of Antibody Purification. J. Chromatogr. B 848, 2007: 48–63.

2 Shukla AA, et al. Downstream Process of Monoclonal Antibodies: Application of Platform Approaches. J. Chromatogr. B 848, 2007: 28–39.

3 Shukla AA, et al. Strategies to Address Aggregation During Protein A Chromatography. BioProcess Int. 3(5) 2005: 36–45.

4 Brorson K, et al. Bracketed Generic Inactivation of Rodent Retroviruses By Low pH Treatment for Monoclonal Antibodies and Recombinant Proteins. Biotechnol. Bioeng. 82(3) 2003: 321–329.

5 Shukla AA, Thommes J. Recent Advances in Large-Scale Production of Monoclonal Antibodies and Related Proteins. Trends Biotechnol. 28(5) 2010: 253–261.

6 Kandula S, et al. Design of a Filter Train for Precipitate Removal in Monoclonal Antibody Downstream Processing. Biotechnol. Appl. Bioc. 54, 2009: 149–155.

7 Yavorsky D, et al. The Clarification of Bioreactor Cell Cultures for Biopharmaceuticals. Pharmaceut. Technol. March 2003: 62–76.

8 Hogwood C, et al. The Dynamics of the CHO Host Cell Protein Profile During Clarification and Protein A Capture in a Platform Antibody Purification Process. Biotechnol. Bioeng. 110(1) 2013: 240–251.

9 Viresolve Prefilter®: Extractables Characterization. EMD Millipore: Billerica, MA, 2012.

10 Yigzaw Y, et al. Exploitation of the Adsorptive Properties of Depth Filters for Host Cell Protein Removal During Monoclonal Antibody Production. Biotechnol. Progr. 22, 2006: 288–296.

11 Haverstock R, et al. CHOP Removal By Depth Filtration Post- Protein A Capture. Abstr. Pap. Am. Chem. Soc. 2010.

12 Zhou JX, et al. Viral Clearance Using Disposable Systems in Monoclonal Antibody Commercial Downstream Processing. Biotechnol. Bioeng. 100(3) 2008: 488–496.

13 Tipton B, et al. Retrovirus and Parvovirus Clearance from an Affinity Column Product Using Adsorptive Depth Filtration. BioPharm Int. September 2002: 43–50.

14 Iskra T, et al. The Effect of Protein A Cycle Number on the Performance and Lifetime of an Anion Exchange Polishing Step. Biotechnol. Bioeng. 110(4) 2013: 1142–1152.

15 Millistak+® Pod Disposable Depth Filter Performance Guide. EMD Millipore: Billerica, MA, 2013.

Corresponding author John Schreffler, PhD, is a group leader, Tom Klimek is a process scientist, Pam Maisey is a researcher, Xun Zuo is a group leader, and Eric Routhier is a director at Eisai, 210 Welsh Pool Road, Exton, PA 19341; 1-610-423-6557; jschreffler@morphotek.com. Corresponding author Matthew Bailley is a process development scientist, Peter Agneta is an account manager, and Michael Felo is a single-use product manager at EMD Millipore, 900 Middlesex Turnpike, Billerica, MA; 1-610-299-5171; matthew.bailley@emdmillipore.com. W. Erick Wiltsie is an associate scientist at GlaxoSmithKline.

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Figure 1: Idealized possible unit structure of sodium carboxymethylcellulose (CMC) with a DS of 1.0; the DS would be 3.0 if all three hydroxyl groups on anhydroglucose unit were substituted. A DS of 3.0 is the theoretical maximum for CMC.

Carboxymethylcellulose sodium (CMC), is widely used as an excipient in oral, topical, and parenteral pharmaceutical formulations. It increases viscosity (1–3), serves as a suspension aid (4), and stabilizes emulsions (5). More recently, applications for CMC in formulations that facilitate improved delivery of cytotoxic drugs and biologics have been evaluated (6, 7).

CMC is manufactured in a broad range of viscosities, with grades typically classified as low, medium, or high viscosity. CMC grades can be divided further based on their degree of substitution (DS), which is defined as the average number of hydroxyl groups substituted per anhydroglucose unit (Figure 1). Together, DS and the extent to which carboxymethyl substituents cluster determine functional properties of CMC (e.g., its aqueous solubility). Thus, CMC offers good water solubility above DS 0.6; at a lower DS (e.g., 0.2), CMC retains the fibrous character of its starting material and is insoluble in water (8).

During the manufacture of parenterals, both finished product and precursory process fluids that are labile to gamma irradiation or heat are protected from microbial contamination by filtration. A sterile filtrate typically can be achieved using a 0.2-μm–rated sterilizing-grade filter that has undergone generic validation by its manufacturer with further support by a user’s process-specific validation.

According to their physical– chemical properties, process fluids show varied filterability (filtration behavior in terms of throughput, as a factor of flow rate and filter membrane capacity to blockage). Viscosity enhancers such as CMC can limit the rate of filtration and incur early filter blockage, impacting upon the the practicality and economy of filter use. In some cases, premature filter blockage and increased processing time associated with filtration of CMC-containing solutions has led to concerns over the practicality and economy of using a sterilizing-grade filter for them at all.

Because of those challenges, opportunities for optimizing the filtration of CMC-based solutions are needed. Here we report on a collaboration between Pall Corporation and Ashland Specialty Ingredients to investigate some factors that can affect the filterability of CMC. Ultimately we seek to provide useful data that can help companies engaged in filtration of CMC-based solutions to make informed choices of filters and CMC grade.

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Table 1: CMC types tested in this study

Materials and Methods
Tables 1 and 2 summarize the materials used in this study. All types of CMC came from Ashland Specialty Ingredients (Wilmington, DE). Filters from Pall Corporation (Basel, Switzerland) were tested as received. Figure 2 shows the experimental setup.

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Table 2: Filters tested in this study (asymmetric layers made to the same specifications)

Preparation of Solutions: CMC particles tend to agglomerate (lump together) when first added to water. For good dissolution, we added CMC to a vortex of vigorously agitated water in preparing test solutions based on the dry mass of CMC (w/w). The rate of addition was slow enough for particles to separate and their surfaces to become individually wetted, but it was fast enough to minimize viscosity buildup of the aqueous phase while the gum was added. Dissolution was complete after an hour of vigorous stirring.

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Figure 2: Experimental setup used for filterability testin

Determination of Viscosity: With test solutions prepared as above, we measured viscosity using a TDVII-LV viscometer from Brookfield Engineering attached by spindle 61 at 60 rpm. Samples were equilibrated at 20 °C before viscosity testing. We calculated the average of three measurements for each solution.

Filterability Testing: Most tests used sterilizing-grade filters without prefiltration. Under those circumstances, 47-mm discs of sterilizing-grade filter membrane were installed in a stainless steel filter housing (Table 2). The housing inlet was connected to an upstream pressure vessel containing a CMC solution to be filtered. The solution passed through the filter by force of air pressure delivered from upstream of the pressure vessel. And filtrate was deposited in a vessel on a balance linked to a system that recorded the change in weight of that collection vessel over time.

When a prefilter was used ahead of a sterilizing-grade filter, the solution to be tested first passed through the prefilter under pressure as above. The resulting filtrate was collected before a second filtration through the final sterilizing-grade filter membrane performed similarly.

All tests were performed at room temperature and at 2-bar constant pressure after ramping up from a starting pressure of 0.5 bar. We terminated each test either when blockage occurred or when sufficient data were collected to indicate that extrapolation of available data would be reliable.

Results and Discussion
Comparison of 0.2-µm–Rated Sterilizing-Grade Filters: We initially subjected Blanose 7M31CF CMC cellulose gum to filterability testing with three different types of 0.2-μm rated, dual-membrane–layer, sterilizing-grade filters differentiated by membrane layer construction and pairing. Table 2 provides information on the construction of each tested filter: Supor EX-grade ECV, Fluorodyne EX-grade EDF, and Supor-grade EBV filter cartridges.

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Figure 3: Comparing filtration throughput performance of Supor EX–grade ECV, Fluorodyne EX–grade EDF, and Supor-grade EBV sterilizing filters with Blanose CMC-grade cellulose gum, 7M31CF at 0.4% concentration

Figure 3 shows the filtration performance of each type of filter membranes (47-mm disc format). We found that the Supor EX-grade ECV filter performed better than both the Fluorodyne EX-grade EDF and Supor-grade EBV filters both in total throughput and filtration speed.

Results for the Fluorodyne EX-grade EDF filter and Supor EX-grade ECV filter reveal how two double-layer sterilizing-grade filters (both using an asymmetric PES prefilter layer of the same specification, but with differing downstream membrane layers) can perform quite differently under the same test conditions. Based on its superior performance, we chose the Supor EX-grade ECV filter as our reference membrane for subsequent investigations into the influence of different characteristics of CMC solutions on filterability.

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Figure 4: Throughput performance of Supor EX–grade ECV filter with Blanose 7M31CF CMC cellulose gum in water

Effect of Concentration and Viscosity: Viscosity of aqueous CMC solutions increases rapidly with CMC concentration. Figure 4 illustrates the filterability-testing results for different concentrations of Blanose 7M31CF CMC cellulose gum in water through a Supor EX-grade ECV filter. Test-solution viscosities were 29.7 mPa•s (0.4 % w/w), 55.4 mPa•s (0.6 % w/w), and 103.8 mPa•s (0.8 % w/w). Observe the rank order between concentration and filterability. Filterability changes significantly depending on concentration (and thus, viscosity) of a CMC solution: The lower the CMC concentration, the better the filterability.

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Figure 5: Throughput performance of Supor EX–grade ECV filter using different Blanose CMC solutions with degrees of substitution (DS) in the range of 0.7–1.2; all solutions were adjusted in concentration to have similar viscosity of 40–60 mPa•s.

DS Effect: Figure 5 shows no significant differences between the filterability of DS 0.7 and DS 0.9 solutions, but a solution with a DS of 1.2 showed a higher tendency for early filter clogging. Blanose CMC cellulose gum comes with three different degrees of substitution: 0.7, 0.9, and 1.2. The most widely used types have a DS of 0.7, or an average of 7 carboxymethyl groups per 10 anhydroglucose units. Higher DS grades give improved compatibility with other soluble components such as salts and nonsolvents due to the ease of hydrogen-bond formation between adjacent chains. The higher the degree of substitution, the more rapidly CMC dissolves. However, depending on the DS grade, different stages of solvatization are reached; correspondingly, higher formation of macrogels and microgels occurs with higher DS (10).

Because of the improved solubility from an increase in DS, we expected that Blanose CMC solutions with a higher degree of DS would have less tendency toward filter clogging than lower-DS solutions. But Figure 5 reveals that our hypothesis — that CMC solutions with a DS of 1.2 would filter more easily than their DS 0.7 counterpart — cannot be confirmed. Results indicate reduced throughput with CMC solutions that have a higher degree of substitution. The effect is not related to viscosity because all CMC solutions were adjusted in concentration to have approximately similar viscosity of 40–60 mPa•s. However, we believe that the reduced filterability of the DS 1.2 solution is attributable to the more extensive formation of microgels at higher DS levels and their more pronounced effect on filterability. Our prefiltration study reports (see below) may help to support these findings.

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Figure 6: Effects of prefiltration on throughput of Supor EX–grade ECV filter membrane with Blanose CMC cellulose gum, DS 0.7 and DS 1.2

Effects of Prefilter/Filter Construction: Prefilters often are used to improve total throughput and flow performance of sterilizing-grade filters. Figure 6 shows the effect of a 2-μm–rated prefilter on throughput performance of a sterilizing-grade filter for CMC solutions with two different degrees of substitution. We kept viscosities of the two solutions (with higher and lower DS) in the same range by adjusting the CMC concentration accordingly (40–60 mPa•s). For the higher degree of substitution (DS 1.2), prefiltration has almost no effect. However, for the lower degree of substitution (DS 0.7), prefilter use remarkably enhances the sterilizing-grade filter’s performance with respect to volume process and filtration speed.

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Figure 7: Effects of sterilizing-grade filter membrane construction on filter throughput using Blanose CMC solutions with different degree of substitution (DS)

We also observed improved filterability of lower-substituted CMC when testing the filterability of CMC solutions with differing degrees of substitution through two sterilizing-grade filters: Supor EX ECV and Supor EBV (Figure 7).

The relationship between filterability of a DS 0.7 CMC solution through Supor EBV filters and Supor EX ECV filters is comparable with that of the filterability of a DS 0.7 CMC solution through a single-stage Supor ECV filter and through the same sterilizing-grade filter following prefiltration (Figure 6).

A Supor EX-grade ECV filter consists of an asymmetric polyethersulfone (PES) membrane layer and a symmetric PES membrane layer; a Supor EBV filter consists of two symmetric PES membrane layers with different retention ratings and a coarser upstream membrane. The asymmetric membrane layer in Supor ECV filters enhances filterability performance with lower-substituted CMC solutions (DS 0.7) but has less effect on higher-substituted CMC solution (DS 1.2).

We can explain the behavior observed in both scenarios by considering the difference in the nature of lower-DS CMC solutions compared with those of higher DS, which are assumed to form more microgels. Throughput performance of a sterilizing-grade filter — either with a built-in prefiltration membrane layer or a separate prefilter — appears to be maintained to a greater extent for lower-DS CMC solutions. At the higher DS of 1.2, behavior exhibited by increased microgel formation appears to reduce the effect of prefiltration.

Challenging Assumptions
Filterability of CMC-containing solutions is affected by a number of variables. Our study addresses filtration more specifically in terms of CMC concentration and degree of substitution as well as sterilizing-grade filter membrane structure and the effect of prefiltration. We observed that the CMC property that is likely to have the most significant influence on filtration was degree of substitution. Best throughputs came with 0.7-DS and 0.9-DS CMC grades, whereas CMC with a DS of 1.2 corresponded with significantly reduced filter-throughput performance. That drop in throughput across all types of filters and filter combinations may be attributable to filter clogging caused by increased formation of microgels at the highest degree of substitution. So using CMC with a DS of 0.7 could offer better opportunities for filter optimization than using CMC with a DS of 1.2.

Concerning different filtration types and filter combinations, use of a prefilter can improve performance of a sterilizing-grade filter downstream for DS 0.7 fluids. Process optimization is otherwise achievable with a sterilizing-grade filter that uses an asymmetric membrane layer and has a high flow rate. We found clearly differentiated filtration performance between two double-layer sterilizing-grade filters with the same asymmetric upstream membrane layer but nonmatching downstream membrane layers. Our observation contraindicates any assumption that all sterilizing-grade filters using an asymmetric prefilter membrane will perform comparably. Wherever possible, filterability testing should be performed on more than one grade of sterilizing-grade filter.

References
1 Shah NH, et al. Carboxymethylcellulose: Effect of Degree of Polymerization and Substitution on Tablet Disintegration and Dissolution. J. Pharmaceut. Sci. 70(6) 1981: 611–613.

2 Wahab A, et al. Formulation and Evaluation of Controlled Release Matrices of Ketoprofen and Influence of Different Co-Excipients on the Release Mechanism. Die Pharmazie 66(9) 2011: 677–683.

3 Khan KA, Rhodes CT. Evaluation of Different Viscosity Grades of Sodium Carboxymethylcellulose As Tablet Disintegrants. Pharm. Acta. Helvetica 50(4) 1975: 99–102.

4 Hussain MA, et al. Injectable Suspensions for Prolonged Release Nalbuphine. Drug Dev. Indust. Pharm. 17(1) 1991: 67–76.

5 Adeyeye MC, et al. Viscoelastic Evaluation of Topical Creams Containing Microcrystalline Cellulose/Sodium Carboxymethyl Cellulose As Stabilizer. AAPS PharmSciTech. 3(2) 2002: E8.

6 Ernsting MJ, et al. Synthetic Modification of Carboxymethylcellulose and Use Thereof to Prepare a Nanoparticle Forming Conjugate of Docetaxel for Enhanced Cytotoxicity against Cancer Cells. Bioconjugate Chem. 22(12) 2011: 2474–2486; doi:10.1021/bc200284b.

7 Sun, et al. Cytokine Binding By Polysaccharide–Antibody Conjugates. Mol. Pharmacol. 7(5) 2010: 1769-77; doi:10.1021/mp100150z.

8 Borsa J, Racz I. Carboxymethylcellulose of Fibrous Character, a Survey. Cellul. Chem. Technol. 29(6) 1995: 657–663.

9 Niu CH, Chiu YY. FDA Perspective on Peptide Formulation and Stability Issues. J. Pharmaceut. Sci. 87(11) 1998: 1331–1334.

10 Gruber E. Mikrogelpartikel in Lösungen von Cellulose und Cellulosederivaten. Cellul. Chem. Technol. 13(3) 1979: 259–278.

Barbara Frei-Rutishauser is a scientist at Pall Corporation in Basel, Switzerland. Christian Muehlenfeld, PhD, is a pharmaceutical scientist, and Gernot Warnke is technical service manager for pharmaceutical R&D, both at Ashland Specialty Ingredients in Dusseldorf, Germany. Corresponding author Tom Watson is a global product manager at Pall Corporation, 5 Harbourgate Business Park, Southampton Road, Portsmouth, UK PO6 4BQ; 44-23-92338185; tom_watson@ europe.pall.com. Supor, Fluorodyne, and Ultipor are trademarks of Pall Corporation.

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Figure 4: Description of Viresolve Prefilter (left) and Viresolve Pro Shield (right) adsorptive prefilters

Size-exclusion–based parvovirus filtration is an important step toward drug product safety in biopharmaceutical production. However, once a virus filter is in place, and the required virus safety is ensured, less attention typically is paid to its optimization within the process. That might seem odd given that virus filtration can be one of the more expensive downstream processing steps ($/g protein processed). Most likely, the lack of attention can be attributed to aggressive timelines, limited process development resources, and the virus filter’s inability to separate drug-product intermediates from host-cell protein and DNA impurities. Most biopharmaceutical process development resources are dedicated to the primary goal of achieving the required drug-product purity. For sound reasons, most of the process-optimization effort goes into critical chromatographic purification unit operations. However, some virus-filter vendors offer significant guidance and extensive hands-on help, which can help biopharmaceutical manufacturers achieve both the highest level of virus safety and virus-filtration cost optimization.

The difficulty of virus-filter separations is underappreciated. Such operations require 100% passage of a ~10-nm monomeric drug-product intermediate (1, 2) and ≥4 log removal of ~20-nm viruses. Furthermore, as drug-product intermediate concentrations increase at the virus-filtration step, and other constraints such as processing-time targets decrease, more process development attention can help optimize this step by making it more efficient and lowering overall cost.

As drug-product concentrations go up, aggregate levels in the intermediate feeds tend to increase as well. That affects virus filters in two ways: First, even without increased aggregates, more monomeric proteins trying to cross the membrane at the same time lowers its starting flux (a polarization effect); second, increased aggregates (dimers, trimers, and so on) can plug filter pores and reduce volumetric throughput (a fouling effect). MilliporeSigma recommends use of in-line adsorptive prefilters to remove foulants (presumably aggregates) and prevent fouling of virus filters (3, 4). An adsorptive-prefilter strategy allows for both optimal flux and volumetric throughput (L/m²). Here we describe optimization efforts using a Viresolve Pro filter coupled in-line with an adsorptive prefilter for a monoclonal antibody (MAb) at 9–13 g/L.

Optimization Parameters (5): Optimization of parvovirus filtration can include changes to pH and/or conductivity from values existing at the preceding chromatographic step, which can reduce aggregate levels in a process stream. Optimization also can include in-line adsorptive prefilters that remove aggregates present in the feed coming directly from the upstream unit operation. In some cases with adsorptive prefilters in place, pH and/or conductivity changes also can be used to optimize the foulant-binding affinity of that prefilter, thereby optimizing virus-filter throughput (3).

Another potential optimization parameter is placement of the virus filter within the process train (following either the second or third column). For particular drug-product intermediates, one placement sometimes works better than another. Virus-filter optimization also can be achieved with higher feed pressures or flows with a higher starting flux (L/m²/h). The fouling profile will be the same — describing a reduction of permeability in L/m²/h/psi with throughput (L/m²) — but more feed can be processed in a fixed period, making the process more productive and time efficient (5).

The strategy we use here strictly focuses on optimization after the third column and running at 30-psi feed pressure. Viresolve Pro filters can run with ≤50-psi feed pressure, but some manufacturing plants limit line pressures to lower levels for safety.

Materials and Methods
The MAb feeds we used were three different third-column pools between 9 and 13 g/L at pH 5.0 and 15 mS/cm conductivity. The first feed (Feed 1) came from multiple elution pools at 9.7 g/L that were generated using smallscale third columns processing a secondcolumn feed from a pilot-scale run. Feeds 2 and 3 were elution pools that came directly from two different thirdcolumn pilot runs (both at 12.4 g/L). All feeds were unfrozen and zero to five days old, stored cold until use, then warmed up to room temperature just before virus filtration. We pH-adjusted some feeds at small scale by placing a pH probe in the feed on a stir plate, making small additions of a buffer concentrate and gently mixing the feed using a magnetic stir bar until its pH reached a target value (5.5–7.0). With pH adjustment, conductivities remained about the same. Before the virus-filter runs, these feeds were 0.2-µm vacuum filtered. The pilot-scale run used similar mixing and buffer addition methods for pH adjustment (in a tank with a top-fed mixer and pump for buffer addition) suited for the larger ~60-L scale.

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Figure 1: Laboratory-scale experimental setup

Laboratory Scale: Each Viresolve Prefilter (catalog #SSPVA40NB9) with 5 cm² of filtration area was wetted and flushed at ~20-psi constant feed pressure for about 10 minutes. After starting slow, the flow increases with processed volume, reaching a final wetted flow rate of ~6 mL/min for each 5-cm² filter. For the wetting and flushing, we used ~45 mL of water per 5-cm² prefilter. After that initial water wet and flush, we flushed buffer through the filter(s) at 20-psi constant feed pressure for four minutes with a flow rate of ~6 mL/min for each prefilter, using about 24 mL per 5-cm² filter. Following buffer equilibration, we assembled the prefilter(s) upstream in-line of a previously prepared Viresolve Pro filter. We used the same procedure for the larger 23-cm² prefilter (#MA1HC23CL3) after venting it of all air with proportionally larger flows and volumes of water and buffer for the larger-area prefilters.

With 3.1 cm² of filtration area, each Viresolve Pro Shield (#VPMSKITNB9) was vented of all air and then wetted and flushed at ~20-psi constant feed pressure for about five minutes. The flow rate was ~15 mL/min. For wetting and flushing, we used ~75 mL of water per 3.1 cm², preparing either one or two in parallel. After the initial water wet and flush, we flushed buffer through the shield(s) at 20-psi constant feed pressure for two minutes. The buffer flow rate was ~30 mL/min for each shield. Buffer flushing used ~60 mL per 3.1-cm² shield. Following buffer equilibration, we assembled the shield(s) upstream in-line of a previously prepared Viresolve Pro filter.

With 3.1 cm² of filtration area, each Viresolve Pro filter (#VPMCPDKNB9) was vented of air and then wetted and flushed at ~50-psi constant feed pressure for about 10 minutes with a flow rate of ~3.5–3.8 mL/min. For wetting and flushing, we used ~37 mL of water. After that, we passed buffer through at 50 psi for four minutes (15 mL total). After that equilibration, we assembled the Viresolve Pro filter in-line downstream of the adsorptive prefilter(s) and ran it with buffer at 30 psi for three minutes before load processing. Then a MAb load was processed at 30-psi constant feed pressure, with cumulative filtrate weight tracked over time. We processed 31–310 mL for each laboratory-scale run.

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Figure 2: Side-by-side laboratory and pilot-scale experimental setups

Pilot Scale: The pilot-scale run used two 0.11-m² Viresolve Prefilters in parallel (#MSPV01FS1), both vented of all air. We flushed them with water using a peristaltic pump at ~2.2 L/min or 600 LMH for about 11 minutes (with 24 L or 109 L/m² total flushed). Feed pressure started at 11 psi and dropped at constant flow to ~6 psi as the filters wetted out. Using the same pump setting, buffer at pH 6.68 and 15.39 mS/cm conductivity was flushed for ~4.5 minutes (12 L total or 55 L/m² at ~2.6 L/min and ~5 psi). Then we assembled the prefilters upstream in-line of a previously prepared Viresolve Pro filter.

The pilot-scale run used a single 0.07m² Viresolve Pro Modus 1.2 device (#VPMD102NB1) vented of all air. We wetted and flushed it with water at a pump speed generating ~30 psi at the filter inlet for about five minutes. Exiting filtrate was collected in 4-L graduated cylinders for tracking cumulative filtrate volume over time. At a flow rate of ~0.9 L/min, we flushed the filter with ~5 L of water. Following the initial water wet and flush, we passed ~2.2 L of buffer through for 2.5 minutes at the same 30-psi feed pressure and 0.9 L/min flow rate. Then we connected this device downstream of the two previously prepared 0.11-m² Viresolve Prefilters.

We set the feed pump speed to generate ~30 psi at the virus filter inlet. With a steady flow achieved through these filters and ~2.2 L of buffer passed through, we could process the MAb load (12.4 g/L) at 30-psi feed pressure. Again, cumulative filtrate weight was tracked over time: ~57 L processed in 2.6 hours (0.5-L/min starting load flow, 0.33-L/min ending load flow). After load processing, we buffer-flushed all feed from the filters to achieve ~100% drug-product yield (10 L of buffer total, flow starting at 0.33 L/min and ending at 0.43 L/min). Following that buffer flush, we flushed the virus filter with water and ran a postuse integrity test using an automatic integrity tester. It passed the test, with a measured diffusion at 50 psi of 1.86 cc/min < 2.7 cc/min.

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Figure 3: Four graphs show laboratory-scale Viresolve Pro device by itself (with no adsorptive prefilter) across a pH range (5.0 with Feed 1; 5.5, 6.5, and 7.0 with Feed 2)

Results and Discussion
Virus Filter Alone: Our approach included both the use of an adsorptive prefilter and a pH change to significantly increase the efficiency and reduce the cost of a virus-filtration step. Throughput with no adsorptive prefilter at the unadjusted pH of 5.0 was only 100 L/m² (Figure 3), which remained the same across a range of pH adjustments (5.0–7.0). So for this particular MAb, the aggregate levels did not lower significantly with pH adjustment. In fact, testing showed that the aggregate levels increased slightly with higher pH levels (data not shown). That slight increase did not further reduce the capacity of the filter alone, however. We found no difference between the feed lots tested (Feed 1 and Feed 2). By contrast, adding an adsorptive prefilter and a pH change to 6.7 increased the achievable throughput to >900 L/m² (Figures 7–9).

Adsorptive Prefilter Options: To be rigorous, we tried two types of adsorptive prefilters in this optimization project: a sterilizing-grade membrane with cation-exchange chemistry and a diatomaceous-earth–containing depth filter. Both prefilters remove foulants by adsorption, the former by ion exchange and the latter by mixed-mode and hydrophobic binding. Either filter can be used effectively to protect a virus filter and reduce costs (3, 4). The Viresolve Pro Shield filter has the benefit of the same module design as the Viresolve Pro device (can be used on the same holder), and as a membrane-based filter it has a relatively low extractables level. It works effectively over a specific range of pH and conductivity compared with the Viresolve Prefilter device. The Viresolve Prefilter requires a separate holder and presents a relatively higher extractables level (Figure 4) (6, 7).

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Figure 5: Viresolve Prefilter (left) and Viresolve Pro Shield (right) performance maps demonstrate their ability to protect a Viresolve Pro filter as a function of feed pH and conductivity, with heatshocked immunoglobulin G (IgG) used as a mock feed stream.

Pros and Cons: MilliporeSigma R&D has conducted experiments with a surrogate feed stream containing high aggregate levels, which provides some guidance regarding optimal pH and conductivity ranges for both the Viresolve Prefilter device and the Viresolve Pro Shield filter to protect a Viresolve Pro filter. At pH levels of 5.0–7.0, the Viresolve Pro Shield’s ability to remove aggregates and protect a virus filter drops off at conductivities in the range of 14–18 mS/cm for the mock feed stream shown (Figure 5).

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Figure 6: Four graphs show Viresolve Pro Shield and Viresolve Pro filters with 1:1 and 2:1 area ratio as a function of pH (5.0 with Feed 1; 5.5, 6.5, and 7.0 with Feed 2).

Viresolve Pro Shield with Viresolve Pro Virus Filter: Because the MAb feed was at pH 5.0 and ~15 mS/cm, we tried the Viresolve Pro Shield filter both at the standard 1:1 (3.1 cm²) and a doubled binding-site 2:1 (6.2 to 3.1 cm²) adsorptive prefilter to virus filter area ratio, both over a pH range (5.0–7.0). Performance increased insignificantly to 100–200 L/m² for all pH levels and both area ratios (Figure 6). That lack of effectiveness was probably attributable to the higher conductivity, which can interfere with an ion-exchange mechanism of aggregate removal. We saw no difference between Feeds 1 and 2. A reduction in feed conductivity would require dilution. We did not pursue that approach because of virus-filter feed-tank volume limitations and the higher volumes that would need to be processed for a downstream ultrafiltration/diafiltration (UF/DF) step.

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Figure 7: Four graphs show Viresolve Prefilter and Viresolve Pro filters with area ratios of 1.6:1.0 cm² and 7.4:1.0 cm² as a function of pH (5.0 with Feed 1; 5.5, 6.5, and 7.0 with Feed 2).

Viresolve Prefilter with Viresolve Pro Virus Filter: The change from the virus filter alone to including a 7.4-cm² to 1.0-cm² area ratio, in-line prefilter at pH 5.0–5.5 increased the throughput from 100 L/m² to >400 L/m², significantly reducing process cost (Figure 7). However, that area ratio is impractical because it requires an unmanageable number of prefilters (exceeding the maximum capacity of the manufacturing plant’s prefiltration holder).

We further optimized this process by changing pH from 5.5 to 6.5 and 7.0. As pH increased, the affinity of the aggregates for the prefilter binding sites increased to the point at which an area ratio of 1.6 cm² (prefilter) to 1.0 cm² (virus filter) was all that was needed to achieve 900 L/m² (Figure 7, bottom-right panel) (3). Note that both area ratios showed the same flat fouling profile at pH 7.0, which means that the affinity of the foulant for the prefilter media is higher at pH 7.0 than at pH 5.5, so the extra prefilter area would not be needed.

Note that we used different feed lots. The first feed (Feed 1) at lower concentration (9.7 g/L) was tested only at pH 5.0. We tested the second feed lot (Feed 2) over a broader range of pH levels (5.5–7.0). Feed 2 was more difficult to filter than Feed 1 probably because of its higher concentration (12.4 g/L) and/or age (≤5 days). We believed that to be the cause of the difference in the 7.4:1.0 area ratio runs shown in the two upper panels of Figure 7.

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Figure 8: Area ratio of 1.6 Viresolve Prefilter to 1.0 (Viresolve Pro) filters as a function of pH (6.5 to 7.0 in increments of 0.1 pH) with Feed 3

Fine-Tuning with pH Optimization: Manufacturing-plant constraints were such that the volume of the virus filter load at pH 5 and added volume for pH adjustment up to 7.0 would exceed existing tank volumes. Combined with a desire to keep the pH further away from the MAb pI (isoelectric point), that situation led to more optimization experiments across pH 6.5–7.0 (Figure 8). We saw no distinct difference across that range, so we targeted pH 6.8 for the first Viresolve Prefilter–Viresolve Pro pilot run. We attributed the difference in performance between Feeds 2 and 3 at pH 6.5 using a 1.6:1.0 area ratio (lower left graphs, Figure 7 and Figure 8) to the freshness of Feed 3 (hours old rather than days). Feed 2 at pH 6.5 achieved 400-L/m² with 90% flow decay, whereas Feed 3 at pH 6.5 achieved 1,000 L/m² with 33% flow decay.

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Figure 9: Area ratio of 3.2 (Viresolve Prefilter) to 1.0 (Viresolve Pro) filters with pH = 6.7 for side-by-side laboratory-scale and pilot-scale run using Feed 3

Robustness and Pilot-Run Scale-Up: One additional consideration was the process robustness and safety factor. We used an area ratio of 3.2:1.0 instead of 1.6:1.0, assuming that would cover potential challenges from even more difficult feeds (concentrations >12.4 g/L, age effects with process delays, and so on). We executed a pilot run processing ~57 L of feed adjusted to pH 6.7 and a 3.2:1.0 ratio. An additional side-by-side laboratory-scale run used the same feed material (Figure 9). Both runs together showed very good scalability and easily achieved >800–900 L/m².

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Table 1: Improvement in parvovirus-filter throughput and process efficiency at different stages in process development; VF = virus filter

Worth the Effort
Our data illustrate the significant benefits that can be reaped through increased time and effort put into optimization of a parvovirus-filtration process (Table 1). The beginning throughput achieved with a virus filter alone at initial processing conditions of pH 5.0 was only 100 L/m². After our optimization efforts, the process achieved a throughput of 1,000 L/m². That was accomplished through use of an adsorptive prefilter with a robust area ratio relative to the virus filter and a pH adjustment that maximized the foulant affinity of the adsorptive prefilter. Optimization efforts like those discussed herein can help biopharmaceutical manufacturers run their rigorously developed, robust parvovirus filtration process as cost-effectively as possible, while still maintaining the highest level of virus safety.

References
1 Reth M. Matching Cellular Dimensions with Molecular Sizes. Nat. Immunol. 14(8) 2013: 765–767; doi: 10.1038/ni.2621.

2 Harris LJ, et al. Refined Structure of an Intact IgG2a Monoclonal Antibody. Biochem. 36(7) 1997: 1581–1597.

3 Bolton GR, Spector S, Lacasse D. Increasing the Capacity of Parvovirus-Retentive Membranes: Performance of the Viresolve Prefilter. Biotechnol. Appl. Biochem. 43(1) 2006: 55–63.

4 Brown A, et al. Increasing Parvovirus Filter Throughput of Monoclonal Antibodies Using Ion Exchange Membrane Adsorptive Pre-Filtration. Biotechnol. Bioeng. 106(4) 2010: 627–637.

5 Lit No. RF1013EN00 Rev. B, 11/14 DP SBU-12-07371: Viresolve® Pro Solution Performance Guide. EMD Millipore Corporation: Billerica, MA, 2014.

6 Technical Brief TB4111ENO, Revision A. Viresolve® Prefilter — Extractables Characterization. EMD Millipore: Billerica, MA, August 2012.

7 Gefroh E, et al. Multipronged Approach to Managing Beta-Glucan Contaminants in the Downstream Process: Control of Raw Materials and Filtration with Charge-Modified Nylon 6,6 Membrane Filters. Biotechnol. Prog. 29(3) 2013: 672–680.

8 Siwak M, et al. Process for Removing Protein Aggregates and Virus from a Protein Solution. US PTO #7,118,675. EMD Millipore: Billerica, MA, 10 October 2006.

9 Siwak M, et al. Process for Removing Protein Aggregates and Virus from a Protein Solution. US PTO # 7,465,397. EMD Millipore: Billerica, MA, 16 December 2008.

Jaime De Souza and Ken Scott are biomanufacturing engineers, and corresponding author Paul Genest is a consulting engineer in MilliporeSigma’s BioManufacturing Sciences Network (BSN), 900 Middlesex Turnpike, Billerica, MA 01824; paul.genest@emdmillipore.com. Viresolve is a registered trademark of MilliporeSigma, a division of Merck KGaA.

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Figure 1: Design and layout of Millidisk and Millipak barrier filters

A number of regulatory guidelines recommend preuse integrity testing of critical sterilizing liquid filters for aseptic processing (1–3). Before sterilization, a preuse test will confirm that a filter is installed properly and was not damaged during shipment or handling. Performing a preuse test after sterilization detects damage that may have occurred during the sterilization cycle. Testing after sterilization limits risk, so it is a practice applied based on risk assessment. Because it is perceived to reduce business loss risk, preuse post-sterilization integrity testing (PUPSIT) is a current industry practice especially in manufacturing products that will be marketed in the European Union (EU).

Unfortunately, it can be difficult to perform a PUPSIT without breaching system sterility. A number of methods have been developed for running PUPSIT and performing line conditioning without compromising sterility. Such methods use a flush bag, catch-can/flush bottle, or filter arrangement to create a sterile boundary on the downstream side of the product filter. Some applications use the downstream hold tank as a sterile boundary.

Here we describe a robust and versatile approach to PUPSIT using a self-venting, all-in-one sterile barrier membrane filter from EMD Millipore, the life science business of Merck KGaA, Darmstadt, Germany, which operates as MilliporeSigma in the United States and Canada. We also include filtration line design considerations for implementing barrier filters.

Barrier Filters
Millipak and Millidisk barrier filters are stacked-disc devices that combine hydrophilic and hydrophobic sterilizing-grade Durapore membranes, both on the top and bottom of each disc (Figure 1). Because of that unique combination of different membranes in parallel configuration, the devices can filter condensate, steam, wetting liquid, and gases without compromising the sterility of a steam-sterilized, autoclaved, or gamma-irradiated system.

Barrier filters can be used downstream of sterilizing-grade filters to maintain system sterility. Water can pass through the hydrophilic discs during a flushing sequence; air can pass through the hydrophobic discs during integrity-test and drying sequences. A barrier filter acts as an automatic vent during testing and drying phases. The volume of particle-free water and air that can pass through these filters is unlimited.

Flushing and Testing Critical Product Filters
Although filter rewetting and retesting should remain an optional activity when preparing a product final filter (sterilizing-grade filter) in line, the PDA Technical Report 26 suggests up to three repetitions (3). The number of retests should be considered when sizing for a flush bag. Barrier filters could provide a more versatile solution.

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Figure 2: Examples of sterile boundary designs

Filters that are not wetted efficiently the first time could give false failed test results. If rewetting volume is limited, end users might discard integral filters that only marginally failed because of improper wetting. Doing so could lead to unnecessary quality investigations as well as downtime associated with setting up a system again before use. Using a barrier filter allows for rewetting and retesting with ease. Unlimited volumes of particle-free water can be filtered through them to wet these filters efficiently. Using a flush bag and/or catch-can instead creates a large footprint and could limit rewetting. Figure 2 summarizes the advantages and disadvantages of each sterile boundary method available.

Barrier filters also allow extractables flushing through a sterile boundary to drain. A barrier filter also can be used as a vent in system cooling after steam-in-place (SIP) sterilization and in filter drying after flushing to minimize product dilution.

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Figure 3: Barrier filters in a typical single filtration system (filling-line example)

Filtration Line Configuration and Operation
For critical filter applications such as final product-filling lines, barrier filters can be used to provide a sterile boundary while meeting regulatory requirements for preuse integrity testing. Typically, such filters are installed downstream of a product filter (Figure 3).

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Figure 4: Product filter preparation steps for steamable line, from sterilizing to integrity testing

Figures 4 and 5 illustrate stepwise use of barrier filters to provide a sterile boundary in a stainless steel and a single-use filtration system with a single product filter. Regulators recommend redundant filtration as a risk-mitigation strategy for critical filtration applications. Redundant filtration is a type of serial filtration in which a second product filter is used as a back-up to protect against the possibility of an integrity failure for the primary product filter (3). Each filter must be independently integrity-testable in compliance with the relevant regulations or guidelines. The step-by-step approaches in Figures 4 and 5 can be applied to the second product filter in a redundant filtration system.

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Figure 5: Product filter preparation steps for single-use assemblies, from sterilizing to integrity testing and from drying to process

Sterilizing: The process begins with sterilization of a filtration system by SIP, autoclaving, or gamma-irradiation. If the chosen method of sterilization is SIP, then the barrier filter’s low-point vent on its upstream side will be kept open during the SIP cycle. Condensate, steam, and air will pass through the filter to drain on its outlet. When the cycle ends, the low-point vent will be closed. The system then cools down with application of compressed gas (to maintain positive pressure as well as sterility in the filtrations system).

Wetting: The second step ensures that a product filter is totally wetted with particle-free water for its integrity test. This step also flushes away extractable residues from a sterilized product filter element. To keep a filter train independent, the vent on the product filter is left open initially, and the isolation valve to downstream equipment is closed. The product filter vent is closed after air in its housing has been vented. Water is directed to the drains through the barrier filters. Isolating the flow path to the barrier filters can enhance wetting for increased applied pressure drop through a product filter.

Testing: PUPSIT is either a diffusion or bubble-point test (depending on the product filter). In all cases, pressurized gas is applied on the product filter’s upstream side, which is isolated from the system elements both upstream and downstream. Only the drain line with the barrier filters remains open to allow the free flow of test gas through the hydrophobic portion of their membranes.

Drying: Before product is introduced into the filtration line, the product filter typically is blown down and dried to prevent dilution of the product stream. The associated gas is vented through the barrier filter.

Barrier Filter Integrity Testing: Millipak and Millidisk barrier filters are integrity tested offline using 70/30 isopropyl alcohol (IPA) as the wetting fluid.

Process: Once the integrity of the product filter is confirmed, the sterile filtration process can begin.

After Processing: After the sterile filtration process, product recovery through a sterilizing-grade filter can be achieved through air blow-down with application of a low differential pressure (air or nitrogen) of 5 psi to the filter. Users can apply a buffer chase, but product dilution must be accounted for. At the end of product recovery, sterilizing-grade filters are integrity tested with particle-free, water-based, alcohol (70/30 IPA/water), or product-based integrity test specifications. For particle-free–water-based or alcohol integrity-test specifications, a filter must be flushed adequately with the test liquid to remove residual product before testing. Product-based integrity-test specifications can be developed through support from filter vendors.

Design Considerations
Use of Millipak and Millidisk barrier filters in a filtration line is simple and straightforward (Figures 4 and 5). Similar to all critical applications, important process steps and conditions should be reviewed during system and process design to ensure successful implementation of this application. Verification testing should be performed before implementation of an assembly with barrier filters for product filtration.

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Figure 6: Sterilization considerations for Millidisk and Millipak barrier filters

Sterilization of Filtration System: Sterilization renders a system or equipment free of microorganisms and is a critical step, especially for aseptic manufacturing processes. Filters can be sterilized through SIP for Millidisk format (cartridge filters) and autoclaving or gamma irradiation for Millipak (disposable capsule) filters (Figure 6). Thermocouples, radiation dosimeters, and biological indicators serve as the worst-case positions within a filtration system and assembly for validation of sterilization.

Flushing and Wetting Product Filters: A filtration system may be flushed and wetted to remove extractables after sterilization as well as for product filter integrity testing. The flushing or wetting liquid passes though the barrier filters to a drain. Flushing/wetting conditions are derived from vendor recommendations (5, 6) and can vary among product filters. Inlet pressures on barrier filters during flushing/wetting procedures should not exceed 0.7 bar. If enhanced wetting of a product filter is required, higher static-hold pressure can be implemented across it. However, the downstream section of that product filter (including the barrier filter) should be isolated during such a high-pressure hold step.

Key Verification Point — Efficacy of Product-Filter Wetting: The efficiency of wetting a product filter(s) through barrier filters can be verified by performing product filter integrity testing. Results can be compared with filter specifications and past trending.

Key Verification Point — Gas Flow Rate of Barrier Filters After Flushing/ Wetting Procedure: Ensuring that the hydrophobic membrane in a barrier filter remains dry is critical. Such dryness can be verified in the filter following a flushing/wetting procedure during the qualification phase with a low-pressure bubble-stream test. This includes disconnecting barrier filters from their assembly and determining their gas flow-rate level at 100 mbar (1.5 psi) pressure, then comparing that to the nominal gas flow-rate level of a new filter unit wetted optimally at low pressure for five minutes. Considering the average hydrophobicity level of polyvinylidene fluoride (PVDF) discs, flow rates in the 30–100% range of nominal rate are characteristic of a “breathing” unit.

Key Verification Point — Using Wetting Medium Apart from Water for Injection (WFI): Compatibility and intrusion pressure/wettability for the hydrophobic filters within barrier filters should be verified before the filters are used. Wetting hydrophobic filters can reduce their air-flow capacity, leading to increased pressure drop across the filter assembly during integrity testing or product-filter blow-down. Millipak and Millidisk barrier filters validation guides provide chemical compatibility information summaries (7–9).

Integrity Testing of Product Filter
Filtration assembly designers should include strategies to minimize product hold by reducing piping or tubing length and to ensure maximum product recovery. The strategy for integrity testing product filters also should be well thought-out, especially for redundant filtration assemblies. In 2012, Felo and coworkers at MilliporeSigma provided an in-depth look at how product filters can be integrity tested in a single-use assembly (10). Their strategy can be applied to stainless steel systems as well.

Interference on Product Integrity Testing from Barrier Filters: Barrier filters are placed downstream of a product filter. To verify the absence of interference, the integrity test result of the product filter both with and without the barrier filters can be compared. Those results should fall within 70 mbar for a bubble-point test and 5% for diffusion flow.

Failure Mode Test: To simulate a worst-case scenario (failure-mode test), users can examine how a fully wet barrier filter gas flow rate compromises a product-filter integrity test. Millipak and Millidisk barrier filters can be fully wetted by flushing with WFI at 3 bar.

Adaptation of Troubleshooting Decision Tree: If a product filter fails its integrity test, users can apply a troubleshooting decision tree such as the example given in PDA’s Technical Report #26 (3).

Drying of Filtration System
To minimize product dilution or contact of product with the wetting liquid (either buffer or water) before filtration, the assembly may be blown down to remove wetting liquid. The current industry practice of blowing down a filtration system ranges from 30 minutes to three hours.

Duration of Drying: Exact drying times should be verified on site and determined during qualification by weight and visual checks. The same time taken to reach the “dry weight” of the assembly will be required for drying the assembly during operation.

Acceptable Applied Pressures: Typical pressures applied for drying filtration assemblies are 0.5 bar higher than the bubble-point pressure of a product filter. Such pressures should not exceed the maximum allowable pressure of the “weakest link” in an assembly. That might be silicone tubing, a connector, or a barrier filter (4.1 bar for Millipak and Millidisk formats), for example. If the required pressure is greater than what the weakest link allows, then blow-down pressure should be reduced, and an extended drying time can be applied.

Absence of Air-Flow Interference: Restriction of air flow through fittings, connectors, tubing, or piping used in an assembly should be minimized. As a product filter dries, the air-flow rate will increase and pressure drop across the product filter will decrease.

Integrity Testing of Barrier Filters
Barrier filters are integrity tested offline with 70/30 IPA/water as a wetting medium and bubble-point test specification of ≥1,280 mbar (18.5 psi). These filters can be wetted by dynamic flushing or static-soak methods. The wetting procedure of barrier filters can be found in a technical guide (5). A 15-minute static soak can be applied to either Millipak or Millidisk barrier filters.

For critical product applications in which resources are readily available, a barrier filter should be integrity tested after the product filter has passed integrity but before product filtration. This minimizes the risk of reprocessing product because of a poor installation or nonintegral barrier filter caused by mishandling. For situations in which resources are limited and product can be reprocessed, barrier filters can be integrity tested after product filtration.

For Best Practices
Barrier filters help enable best practices of aseptic filtration lines for flushing/wetting and preuse integrity testing of product filters. In particular, implementation of Millipak and Millidisk barrier filters is easy and provides flexibility and versatility to the filtration line.

References
1 Annex 1: Manufacture of Sterile Medicinal Products. Volume 4, EU Guidelines to Good Manufacturing Practice Medicinal Products for Human and Veterinary Use. European Commission: Brussels, Belgium, November 2008; http://ec.europa.eu/health/files/eudralex/vol-4/2008_11_25_gmp-an1_en.pdf.

2 CBER/CDER/ORA. Sterile Drug Products Produced By Aseptic Processing: Current Good Manufacturing Practice. US Food and Drug Administration: Rockville, MD, September 2004.

3 Technical Report No. 26: Sterilizing Filtration of Liquids. Parenteral Drug Association: Bethesda, MD, 2008.

4 ASTM Standard F838-83: Standard Test Method for Determining Bacterial Retention of Membrane Filters Utilized for Liquid Filtration. American Society for Testing and Materials: Philadelphia, PA, 1983.

5 P35515 Rev G: Wetting Instructions for Filter Units with Durapore Membrane. EMD Millipore: Billerica, MA, April 2012.

6 RF1510EN00: Hydrophilic Durapore Cartridges and Capsules User Guide. EMD Millipore: Billerica, MA, January 2002.

7 VG026 rev 2: Millidisk Cartridge Filter Units with Hydrophilic Durapore Membrane Validation Guide. EMD Millipore: December 1999.

8 VG033 Rev D: Millipak Disposable Filter Units Validation Guide. EMD Millipore: Billerica, MA, May 2012.

9 VG2000EN00: Millidisk Barrier Filter Validation Guide. EMD Millipore: Billerica, MA, May 2002.

10 Felo M, Oulundsen G, Patil R. SingleUse Redundant Filtration. BioPharm Int. 25(4) 2012: 38–41.

Corresponding author Yanglin Mok, BE, is a senior process engineer and technical manager of the Biomanufacturing Sciences Network, 1 Science Park Road, #02-10/11 The Capricorn, Singapore 117528; 65-6403-5313, fax 65-6403-5322; yanglin.mok@merckgroup.com. Lise Besnard, MSc, is a process development scientist at Sanofi Pasteur, 1541 Avenue Marcel Mérieux, 69280 Marcy l’Etoile, France. Terri Love, BSc, is a biomanufacturing engineer; Guillaume Lesage, MSc, is a biosafety technical consultant; and Priyabrata Pattnaik, PhD, is director of the worldwide vaccine initiative; all with the life-science business of Merck KGaA, Darmstadt, Germany, which operates as MilliporeSigma in the United States and Canada. Millipak, Millidisk, and Durapore are registered trademarks of MilliporeSigma.

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Figure 1: General polysaccharide conjugate vaccine process (UF/DF = ultrafiltration/diafiltration)

Polysaccharide vaccines are essential for protection against infectious diseases, which remain an alarming cause of mortality. The first glycoconjugate vaccine for use in humans — a Haemophilus influenzae type b (Hib) conjugate — was licensed in the United States in 1987. This vaccine successfully reduced the incidence of invasive Hib disease in childhood and led to the further development of conjugate vaccines designed to prevent infection by other encapsulated bacteria (1).

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Table 1: General polysaccharide antigen properties

Polysaccharides are relatively complex carbohydrates made up of many monosaccharides joined together by glycosidic bonds. Bacterial polysaccharides represent a diverse range of macromolecules that include peptidoglycans, lipopolysaccharides, capsules, and exopolysaccharides — compounds whose functions range from structural cell-wall components (e.g., peptidoglycan) to important virulence factors (e.g., Streptococcus pneumoniae, Neisseria meningitidis, and Haemophilus influenzae). Most polysaccharides produced by bacteria are of high molecular weight with acidic isoelectric points (pI). Table 1 shows general properties of polysaccharide antigens.

Polysaccharide antigens consist of repeating epitopes that are not processed by antigen-presenting cells (APCs). These antigens interact directly with B cells, which can induce antibody synthesis in the absence of T cells (known as T-independent antigens). T cells can influence the antibody response to certain polysaccharides, such as the capsular polysaccharide of S. pneumoniae type 3. T-independent responses fail to induce significant and sustained amounts of antibody in children below the age of 18 months (1, 2). In 1929, Avery and Goebel demonstrated that the poor immunogenicity of purified S. pneumoniae type 3 polysaccharide in rabbits could be enhanced by conjugation of the polysaccharide to a protein carrier. That led to the foundation for the development of conjugated polysaccharide vaccines (1).

Modern polysaccharide vaccines are generally conjugated to nontoxic, nonreactogenic carrier proteins or tetanus toxoid (a 150-kD protein from the Gram-positive anaerobic bacteria Clostridium tetani). Alternatively, CRM 197 (a single point-mutated 68-kD protein purified from Cornybacterium diptheriae or as recombinant protein expressed in Eshcherichia coli or Pseudomonas) has been the carrier protein of choice in several vaccines. The selection of a carrier protein can be driven by a number of factors, including availability, price, and chemical characteristics such as stability at certain pHs and adjuvant effect (2).

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Table 2: Licensed carbohydrate-based vaccines (13)

Several polysaccharide-based vaccines are in current development pipelines. Carbohydrate-based vaccines in development are being derived from Streptococcus, Pseudomonas aeruginosa, Salmonella typhi, Shigella dysenteriae, Shigella flexneri, Shigella sonnei, Vibrio cholera, Leishmania species, and others (3). Table 2 shows a selection of carbohydrate vaccines that have been licensed.

There are several other carrier proteins used as conjugation partners for polysaccharide vaccines. Pneumococcal surface protein A (PspA) is one such carrier protein (3). Another is protein D (PD), a 42-kDa surface lipoprotein found in all Haemophilus influenzae, including nontypeable (NT) H. influenzae used as an antigenically active carrier protein in an 11-valent pneumococcal conjugate investigational vaccine (4). Diphtheria toxoid (DT), tetanus toxoid (TT), and CRM197 also have been used as protein carriers in licensed vaccines (5).

Vaccines for N. meningitidis are based on outer membrane vesicles between 20 and 200 nm. Outer membrane vesicles (OMVs) are released spontaneously during growth by multiple Gram-negative bacteria. OMVs have lipopolysaccharides, phospholipids, proteins, RNA/DNA, and peptidoglycan, and they are produced by bacterial fermentation. Centrifugation has been implemented for recovery. Tangential-flow filtration (TFF) cassettes (10–100 kDa) also have been used to separate OMV from small soluble components (6). TFF provides a range of surface antigens that are in native conformation and possess natural properties such as immunogenicity, self-adjuvation, and uptake by immune cells. Such characteristics make them attractive for applications as vaccines against pathogenic bacteria. An OMV-containing meningococcal vaccine (Bexsero from Novartis) was recently approved by US and EU regulatory agencies, and research on its application continues.

Despite growing interest, no established template exists for a vaccine purification platform because of the complexity and diversity of vaccines (7). The manufacturing process of vaccines can be divided into three segments: upstream processing, downstream processing (purification), and formulation (fill–finish operation). Figure 1 illustrates a generic conjugated polysaccharide vaccine production process.

Clarification is an essential operation in the production of biological products because it directly affects yield, product consistency, and reproducibility. The goals of clarification include high yield, product consistency, and reproducibility. Primary clarification removes the majority of large particles, whole cells, and cell debris. This step can be performed by TFF–microfiltration (TFF–MF), centrifugation, or (in some cases) depth filtration. Secondary clarification is used to remove colloids, lipids, DNA–RNA, residual cells, and other particles not removed in a primary clarification process. Secondary filtration typically includes a series of filters to remove progressively smaller particles (8).

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Table 3: Combination of technologies used for clarification of vaccines

Traditionally, a combination of technologies has been used for the clarification of conjugated polysaccharide vaccines, as summarized in Table 3. Because centrifugation can handle high solid loads, it has been the preferred method. Generally, initial cell mass is in the range of 7–30 OD at 590 nm or can reach 5 g/L dry cell weight (9). Centrifugation can concentrate cell mass to about 40% of the initial volume (8). Clarifying filtration can be performed by normal-flow filtration (NFF, also known as dead-end filtration) or TFF (also known as cross-flow filtration). Depth filters contain positively charged material and filter aid that enhance retention of cell debris, colloids, and unwanted negatively charged components (8, 10). Membrane filters retain particles by size exclusion, but they have limited dirt-holding capacity and are more suitable for a secondary clarification step.

Scalability is not a concern with depth filters or membrane filters. TFF is used mostly for primary clarification (microfiltration) with successful cut-offs in the range of 0.1–0.65 µm (preferably with open channel). Linear scalability and reusability of TFF devices significantly reduces costs of consumables in the clarification step (11).

Improper optimization of clarification steps can affect the performance of subsequent downstream unit operations in terms of capacity of filters or life of membranes and resins. With a need for well-characterized vaccines, simplified processes, as well as increased purity, new filtration technologies can handle clarification challenges for more process flexibility, possibility of single-use, and reduced investment costs.

Clarification of Polysaccharide Vaccines
Considerations for Polysaccharide Vaccines Clarification: After fermentation of polysaccharide vaccines, the next step is harvest clarification. For high-packed cell volumes, direct filtration through normal-flow filters is not economically feasible because of low throughput. In most cases, centrifugation is common practice for separation of cell mass. TFF–MF also could be used (12–14). Filtrate from the TFF–MF step can be further clarified using NFF depth filtration train, followed by bioburden reduction by filtration. In some cases, homogenization can be implemented to enhance performance of clarification (15).

Buffer conditions and pH can affect polysaccharide clarification.Hadidi et al. demonstrated the effect of ionic strength and pH on the hydrodynamic radius of the free polysaccharide and conjugated polysaccharide, which then had an impact on retention time during size-exclusion chromatography (SEC) (16). The study also showed that retention time of the pneumococcal polysaccharides increases in SEC columns with increasing ionic strength and decreasing pH because of compaction of the polysaccharides associated with a reduction in intramolecular electrostatic interactions.

Strategy for Polysaccharide Vaccines Clarification
Primary Clarification Step: Centrifugation is the preferred technology for separating high cell mass from fermentation broth. Depending on scale, continuous or batch centrifugation could be used. General centrifugation conditions are 14,000–15,000g for 45–60 minutes (17, 18).

TFF at microfiltration (MF) range can be used as an alternative to centrifugation. The molecular weight of polysaccharides that are large and complex in structure typically ranges from about 500 kDa to over 1,000 kDa. MF membranes (e.g., 0.22 µm, 0.45 µm, and 0.65 µm) are preferred to ensure successful recovery of polysaccharide molecules in permeate because of their large open pore size.

Secondary Clarification Step: The clarity/turbidity of cell-free fermentation broth depends on the specific bacteria, lysis type, individual serotype, and technology used for primary clarification. Turbidity of postcentrifuge centrate could range from about 50 NTU to 1,300 NTU. After primary clarification steps, NFF with depth media (e.g., B1HC, C0HC, F0HC, or X0HC grades of Millistak Pod disposable depth filters from MilliporeSigma) could be used to achieve turbidity <5–10 NTU, suitable feed levels for further purification steps.

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Figure 2: General depth filter performance for clarification of harvest

Volumetric throughputs on the depth filter can range from about 30 L/m² to 500 L/m². Depth filter clarified product fluid could be filtered through a subsequent 0.45-µm bioburden-reduction–grade or 0.22-µm sterilizing-grade membrane. In general, performance of depth filters varies among different polysaccharide vaccines, and values and performance might vary from case to case. Feed can range between 100 NTU and 10,000 NTU. Depth filters in general have shown capacities of 50–300 L/m². In the postprimary clarification, the filtrate turbidity can drop down to <50 NTU. Figure 2 shows performance of several depth filters.

Case Studies
Clarification of Postcentrifuge Centrate of S. pneumoniae Fermentation Broth: The traditional method of clarification is centrifugation, followed by time-consuming, multiple-step depth filtration. Some researchers have made efforts in developing single-step, filtration-based clarification methods for polysaccharide vaccines. In one study, charged depth filters were implemented for clarification of S. pneumoniae serotype 9V fermentation broth, resulting in 10- to 13-fold reduction of turbidity from 130 NTU (data not published). Such one-step clarification operated at flow rate of 50–60 L/m²/h (LMH) resulted in completion of clarification of a 2,000-L batch in about three to four hours. This translated to a volumetric throughput of about 180–200 L/m². Similar filtration performance also was observed with S. pneumoniae serotype 20F, 12F, and 17F.

As a result of this study, the method of centrifugation for clarification of pneumococcal harvest was changed to depth filtration. The ideal way to judge the effect of a change in clarification method is to observe performance of downstream filtration. If filtrate quality improves, downstream filtration performance ideally would improve in terms of capacity (volumetric processing) and can lower differential pressure due to slower plugging.

Clarification of Postcentrifuge Centrate of H. influenzae Fermentation Broth: Centrifugation is generally the preferred choice for primary clarification. But in 2008, Takagi evaluated a two-way approach (19). First, H. influenza B fermentation broth was loaded onto depth filters. Filtration was performed using a train of two charged depth filters. Reduction of feed turbidity was observed from about OD 5 to less than OD 0.1 at 650 nm. Tests were performed at 10–30 LMH, and 300 L were processed in about 3–4 hours with capacities of about 20–30 L/m².

The other approach was to subject the harvest to microfiltration TFF (using two pump-based, permeatecontrolled operation) as an alternative, reusable option to depth filters (20). After this TFF processes the feed, OD of the filtrate decreased from 5 to <0.1. The flux was 10 LMH, with loading of about 30–40 L/m². Area requirement for processing 300 L of volume was 10 m² in three hours (unpublished data).

If possible, single-step TFF is a good alternative to centrifugation and secondary clarification. Scaling-up or scaling down centrifuge is not easy. Depending on specific conditions, TFF cassettes can be used for multiple batches.

Clarification of Salmonella typhi Vi Harvest: Inactivated Salmonella typhi bacteria can be clarified using a two-step process. As an option for primary clarification, a TFF device (0.45-µm cassette) was used to concentrate bacteria cells 7–10-fold and then diafiltered 10 times against 1M NaCl. During cell concentration, some Vi polysaccharides passed through the membrane into the permeate. A significant quantity of Vi polysaccharides is generally retained because of the ionic interaction between the negatively charged Vi polysaccharides and other positively charged components in the broth. A high salt concentration can be used to weaken those interactions — neutralizing the strong negative charge on the Vi polysaccharides (at pH ∼7.0), facilitating passage of Vi polysaccharides through membrane pores, and resulting in increased recovery of Vi polysaccharides (12, 13). Typical volumetric loading for such operation is 45–55 L/m² and the processing can take three to five hours.

As a secondary clarification option, 30-kDa TFF can be used on the permeate pool from first TFF step for concentration (10–15 fold) and diafiltration (8–10 times) against water for injection (WFI). During this step, Vi polysaccharides will remain in the retentate, and low–molecular-weight impurities and excess water would pass through the membrane into the permeate. Diafiltration against water was required to remove excess salt to make negatively charged Vi polysaccharides readily accessible to the positively charged Cetavlon for subsequent precipitation step (12, 13). Typical volumetric loading for such operation is 45–55 L/m², and the processing can take 3–5 hours, resulting in about 100% recovery.

TFF coupled with changes in solubility or ionic interactions facilitate clarification procedures and help reduce contaminant load and size-dependent impurities. This results in high recovery of polysaccharides.

Clarification of Postcentrifuge Centrate of Fermentation Broth By Addition of a Flocculating Agent: Adding flocculating agent to a bioreactor before centrifugation can improve depth-filtration–based clarification. A study conducted in our laboratory evaluated multiple depth filters to select an appropriate secondary clarifying filter. The study found that implementation of a Millistak+ C0HC filter (MilliporeSigma) alone as a secondary clarification filter on centrate reduced turbidity by ∼90% (from feed NTU 100, to 10 NTU). Filtration using a Millistak+ C0HC filter was performed at ∼500 LMH, resulting in a capacity >400 L/m² (unpublished data). With a Millistak+ C0HC filter, a minimal ∆P was observed, suggesting that the centrifuge is removing larger particles, and the Millistak+C0HC filter (with a tighter nominal pore size) is probably removing smaller particles.

Those results indicate that with proper screening of depth filters, a filtration train can be reduced while still achieving a desired throughput and reduction in turbidity. In turn, that may result in smaller footprint and ease of operation. In our study, only one step depth filtration achieved the desired clarification of centrate.

A number of next-generation filtration products are being introduced to enable better, easier, and more robust clarification applications. Kang et al. observed that Clarisolve filters (from MilliporeSigma) can be implemented for better clarification of flocculated monoclonal antibody harvest (14). Those filters are more open, with pore sizes 20–60 µm in range.

In a study on pneumococcal vaccine, Clarisolve 60HS (polypropylene) filters (from MilliporeSigma) were evaluated on a harvest pretreated with celite and cetyltrimethylammonium bromide (CTAB). This study resulted in loading of 50 L/m² and found that performance of Clarisolve filters as primary clarifying filters do not depend on cell viability and cell count. Implementation of Clarisolve filters for pretreated polysaccharide vaccine harvest can omit centrifugation and secondary clarification steps, resulting in a smaller footprint and easier operation.

Next-Generation Clarification Methods
Clarification of polysaccharide vaccines presents several challenges. Typically, filtration processes and filtration trains vary case by case. Because of high cell mass, centrifugation or microfiltration devices typically are preferred for primary clarification. Of late, charged depth filters have shown promise for primary clarification, and membrane filters are proving to be better options for secondary clarification.

Templates for purification of polysaccharide vaccines are being implemented or adopted, and clarification schemes by filtration are delivering high degrees of success due to robustness, ease of scalability, and process economics. Improvement in clarification steps have resulted in higher final yields and purity in vaccine processes. As new clarification products, tools, and solutions are being made available, vaccine developers and producers will continue to be better prepared for efficient and effective clarification processes.

References
1 Goldblatt D. Conjugate Vaccines. Clin. Exp. Immunol. 119, 2000: 1–3. doi: 10.1046/j.1365-2249.2000.01109.x.

2 An SJ, et al. Physicochemical Properties of Salmonella typhi Vi Polysaccharide: Diphtheria Toxoid Conjugate Vaccines Affect Immunogenicity. Vaccine 29, 2011: 7618–7623. doi: 10.1016/j.vaccine.2011.08.019.

3 Astronomo RD, Burton DR. Carbohydrate Vaccines: Developing Sweet Solutions to Sticky Situations? Nat. Rev. Drug Discov. 9(4) 2010: 308–324. doi: 10.1038/nrd3012. PMID 20357803.

4 Forsgren A, et al. Protein D of Haemophilus influenzae: A Protective Nontypeable H. influenzae Antigen and a Carrier for Pneumococcal Conjugate Vaccines. Clin. Infectious Diseases 46, 2008: 726–731. doi: 10.1086/527396.

5 Tontini M, et al. Comparison of CRM197, Diphtheria Toxoid and Tetanus Toxoid as Protein Carriers for Meningococcal Glycoconjugate Vaccines. Vaccine 31(42) 2013: 4827–4833. doi: 10.1016/j.vaccine.2013.07.078.PMID 23965218.

6 van der Pol L, et al. Outer Membrane Vesicles as Platform Vaccine Technology. Biotechnol. J. 10(11) 2015: 1689–1706. doi: 10.1002/biot.201400395.

7 Ball P, et al. 21st Century Vaccine Manufacturing: Examining the Potential of Rapid Analytical Methodologies and Worldwide Supply Chains. BioProcess Int. 7(4) 2009: 18–28.

8 Yavorsky et al. The Clarification of Bioreactor Cell Cultures for Biopharmaceuticals. Pharma. Technol. 2003; 62–76.

9 Fu J, et al. Recent Advances in the Large-Scale Fermentation of Neisseria meningitidis Group B for the Production of an Outer Membrane Protein Complex. Biotechnol. 13(2) 1995: 170–174. PMID 9634759.

10 van Reis R, Zydney A. Membrane Separations in Biotechnology. Curr. Op. Biotechnol. 12(2) 2001: 208–211. doi: 10.1016/S0958-1669(00)00201-9.

11 Besnard L, et al. Clarification of Vaccines: An Overview of Filter-Based Technology Trends and Best Practices. Biotechnol. Adv. 34(1) 2016: 1–13. doi: 10.1016/j.biotechadv.2015.11.005.

12 Kothari S, et al. A Novel Method for Purification of Vi Capsular Polysaccharide Produced By Salmonella enterica Subspecies Enterica Serovar Typhi. Vaccine 31(42) 2013: 4714–4719. doi: 10.1016/j.vaccine.2013.08.037; PMID 23994374.

13 Kothari S, et al. Development of an Efficient and Scalable Method for Processing and Purification of Vi Capsular Polysaccharide. Proc. Vaccinol. 2, 2010: 78–81. doi: 10.1016/j.provac.2010.03.014.

14 Gonçalves VMM, et al. Purification of Capsular Polysaccharide from Streptococcus pneumoniae Serotype 23F By a Procedure Suitable for Scale‐Up. Biotechnol. Appl. Biochem. 37(3) 2003: 283–287. doi: 10.1042/BA20020075.

15 Andre BR, Champluvier BPS, inventors; Glaxosmithkline Biologicals SA, assignee. Method for Producing Virus from Cell Culture Involving Homogenization. WO2010128100A1, 11 November 210.

16 Hadidi M, et al. Effect of Solution Conditions on Characteristics and Size-Exclusion Chromatography of Pneumococcal Polysaccharides and Conjugate Vaccines. Carbohydrate Polymer 152, 2016: 12–18. doi: 10.1016/j.carbpol.2016.06.095.

17 Pisal SS, Reddy CS, Reddy PS, inventors; Serum Institute of India Ltd, assignee. Production of High Yields of Bacterial Vaccines. WO 2014080423A2. 30 May 2014.

18 Peddireddy SR, et al., inventors; Serum Institute of India Ltd, assignee. Bacterial Capsular Polysaccharide Yield Enhancement By Addition of Defoaming Agents. EP2952586A1. 9 December 2015.

19 Takagi M, et al. Purification of Capsular Polysaccharide Produced By Haemophilus influenza Type b Through a Simple, Efficient, and Suitable Method for Scale-Up. J. Ind. Microbiol. Biotechnol. 35(11) 2008: 1217–1222. doi: 10.1007/s10295-008-0428-4.

20 Raghunath B, et al. Best Practices for Optimization and Scale-Up of Microfiltration TFF Processes. Bioprocess J. 11(1) 2012: 30–40; http://dx.doi.org/10.12665/J111.Raghunath.

21 Liu TY, et al. Studies on the Meningococcal Polysaccharides, 1: Composition and Chemical Properties of the Group A Polysaccharide. J. Biol. Chem. 246(9) 1971: 2849–2858.

22 Liu TY, et al. Studies on the Meningococcal Polysaccharides, 2: Composition and Chemical Properties of the Group B and C Polysaccharide. J. Biol. Chem. 246(15) 1971: 4703–4712.

23 Apicella MA, Robinson JA. Physicochemical Properties of Neisseria meningitidis Group C and Y Polysaccharide Antigens. Infect Immun. 2(4) 1970: 392–397.

24 Apicella MA, Robinson JA. Physicochemical Properties of Neisseria meningitidis Group X Polysaccharide Antigen. Infect. Immun. 6(5) 1972: 773–778.

25 Jedrzejas MJ. Pneumococcal Virulence Factors: Structure and Function. Microbiol. Mol. Biol. Rev. 65(2) 2001: 187–207. doi: 10.1128/MMBR.65.2.187–207.2001.

26 Briles DE, et al. Pneumococcal Diversity: Considerations for New Vaccine Strategies with Emphasis on Pneumococcal Surface Protein A (PspA). Clin. Microbiol. Rev. 11(4) 1998: 645–657.

27 Worthington Biochemical Corp. Neuraminidase; www.worthington-biochem.com/NEUP/default.html.

28 Gürtler L. Virology of Human Influenza. Influenza Report 2006. Kamps BS, et al., Eds. Flying Publisher: Paris, France, 2006: 87–91.

29 Shtyrya YA, et al. Influenza Virus Neuraminidase: Structure and Function. Acta Naturae 1(2) 2009: 26–32. PMID22649600.

30 Complete List of Vaccines Licensed for Immunization and Distribution in the US; www.fda.gov/BiologicsBloodVaccines/Vaccines/ApprovedProducts/ucm093833.htm.

31 Perciani CT, et al. Conjugation of Polysaccharide 6B from Streptococcus pneumoniae with Pneumococcal Surface Protein A: PspA Conformation and Its Effect on the Immune Response. Clin. Vaccine Immunol. 20(6) 2013: 858–866. doi: 10.1128/CVI.00754-12.

32 Robinson A, et al. Meningitis Vaccine Manufacturing: Fermentation Harvest Procedures Affect Purification. BioPharm. Int. 24, 2011: s21–s26.

33 Kalbfuss B, et al. Harvesting and Concentration of Human in Influenza A Virus Produced in Serum-Free Mammalian Cell Culture for the Production of Vaccines. Biotechnol. Bioeng. 97(1) 2007: 73–85. doi: 10.1002/bit.21139. PMID 16921531.

34 Macha C, et al. Purification of Streptococcus pneumonie Capsular Polysaccharides Using Aluminium Phosphate and Ethanol. Int. J. Pharmacy Pharm. Sci. 6(2) 2014: 385–387.

35 Hamidi A, Haag D, Beurret MF, Bilt D, inventors. Nederlands Vaccin Instituut, assignee. Process for Producing a Capsular Polysaccharide for Use in Conjugate Vaccines. US patent 2007/0065460 A1. 22 March 2007.

36 Hamidi A, et al. Process Development of a New Haemophilus influenzae Type B Conjugate Vaccine and the Use of Mathematical Modelling to Identify Process Optimization Possibilities. Biotechnol. Prog. 32(3) 2016: 568–580. doi: 10.1002/btpr.2235.

37 Kang Y, et al. Development of a Novel and Efficient Cell Culture Flocculation Process Using a Stimulus Responsive Polymer to Streamline Antibody Purification Processes. Biotechnol. Bioeng. 110(11) 2013: 2928–2937. doi: 10.1002/bit.24969.

Nikhil Shaligram is manger of process development at Merck Life Science Pvt Ltd. (Mumbai, India). Sudeep Kothari is senior research scientist at Vaccine Process Development, Science Division, International Vaccine Institute (Seoul, Korea). Keunhoe Koo is senior process development scientist and Cheon-Ik Park is manager of biomanufacturing science and technology at Merck Ltd. (Seoul, Korea) Elizabeth Goodrich is head of applications engineering at MilliporeSigma Corporation (Billerica, MA). Corresponding author Priyabrata Pattnaik is director and head of biologics operations at Merck Pte. Ltd., 3 International Business Park, #02-01 Nordic European Centre, Singapore 609927; 65-6890-0610; fax 65-6890-6776; priyabrata.pattnaik@merckmillipore.com.

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Figure 1: Right-sizing and rapid multicycling enables small-footprint downstream processing to match high-productivity upstream production (adapted from reference 1).

Filtration membranes are used extensively throughout the biopharmaceutical industry for a range of applications, from coarse filtration to nanofiltration. Advantages of filter technologies include easy scaling, disposability, and (for many membrane filters) rapid and robust performance in a single-pass. The same advantages have been realized with membrane adsorbers.

Chromatography resins are inherently disadvantaged by diffusion limits of the pores in chromatography media. Therefore, resin columns must be significantly oversized to match the performance of high productivity bioreactors. By comparison, membrane adsorbers take advantage of filter performance to deliver highly productive downstream operations. Surface-functionalized membranes from Sartorius Stedim Biotech (Sartobind Q, STIC) and Pall Life Sciences (Mustang Q)typically use anion-exchange groups for MAb polishing operations in negative mode (in which trace impurities are removed without binding the protein of interest). Such flow-through chromatography operations for impurity reduction can enhance purification productivity by over an order of magnitude: from ≤250 g/L capacity (~250 g/L-h productivity for 10 g MAb/L and two minutes residence time representing one hour operation) for chromatography resins to ≤10,000 g/L capacity (~5,000 g/L-h productivity for 10 g MAb/L and six seconds residence time representing two hours operation) for membranes.

Natrix Separations’ 3-D hydrogel membrane columns (NatriFlo HD-Q) demonstrate performance up to 20,000 g/L membrane for MAbs, with impressive host cell protein (HCP) and virus reduction with only six seconds of residence time. Membrane adsorbers are increasingly used in new flexible manufacturing facilities, in which speed of operations and robust performance are essential production criteria. These functional membranes are the first truly disposable, single-use chromatographic media with best-in-class throughput for polishing.

Membrane Chromatography: Advanced Materials
Cation-exchange operations typically show excellent aggregate reduction and HCP removal for chromatography resins. Negative-mode chromatography also can be used for significantly increased productivity. Although traditional membrane adsorbers have limited capacity, next-generation membranes have 10× more capacity than traditional membranes. Their media can be fully used in a single batch by “right-sizing” membrane columns and performing rapid cycling.

Therefore, other chromatography modalities have been developed. Sartobind S (Sartorius Stedim Biotech) and Mustang S (Pall Life Sciences) show good performance for cation-exchange chromatography, but their binding capacity is still limited by low ligand density per milliliter of membrane media. Natrix Separations is developing a salt-tolerant hydrogel membrane (Natrix HD-Sb) that can maintain high capacity and operate in either positive or negative mode (≤90 g/L or ≤ 500 g/L membrane, respectively), with good aggregate clearance.

In addition, membranes with other functional groups are in development for a range of purification operations. These include hydrophobic-interaction chromatography (Sartorius Stedim Biotech) and affinity chromatography (Sartorius Stedim Biotech and Natrix Separations). These new modalities offer full process platforms that promise to match the production rates of upstream bioreactors with appropriate downstream operations for MAb, vaccine, and virotherapy applications.

Single-Use Membrane Columns: Productivity and Flexible Manufacturing
Outputs of single-use bioreactors for commercial MAb production now range from 500 g to 20 kg at harvest. Resin columns (now offered in “single-use” prepacked format) traditionally have been used to handle such protein quantities. However, to maintain good process throughput, again the columns need to be oversized. That can be very costly especially for clinical production, in which the full resin cost is amortized over limited use. In commercial production, an expensive resin’s full useful life can be exploited.

A new paradigm in biomanufacturing involves high-productivity single-use bioreactors and a train of single-use (per batch) membrane chromatography columns that can keep pace with the upstream productivity (Figure 1). Membrane chromatography offers a powerful alternative for improved process throughput, economics, validation, and quality oversight. The strategy includes flow-through membrane adsorbers along with bind/elute membrane columns (e.g., for protein A affinity membranes) that can perform like packed-bed resins for capacity and separation but operate one to two orders of magnitude faster (2). That speed drives productivity (applied practically, not theoretically) to 30× more than resins for bind–elute chromatography.

Membrane columns can reduce operational time, cost, and complexity while maintaining high levels of production output per week for commercial biomanufacturing. A 1,000-L bioreactor culture operating at 5-g/L expression levels and a single membrane chromatography downstream train can produce 4 kg/batch of drug substance (DS). With more than one bioreactor feeding that single right-sized downstream train, production of 400 kg DS per year or more could be achieved in a fully flexible and cost-efficient facility.

Membrane chromatography is coming of age. We believe that it is the future of highly productive, flexible manufacturing for clinical and commercial protein production (3).

References
1 Jacquemart R, et al. A Single-Use Strategy to Enable Manufacturing of Affordable Biologics. Comput. Struct. Biotechnol. J. 14, 2016: 309–318; doi:10.1016/j.csbj.2016.06.0072016.

2 Hou Y, et al. Advective Hydrogel Membrane Chromatography for Monoclonal Antibody Purification in Bioprocessing. Biotechnol. Progr. 31, 2015: 974–982; doi:10.1002/btpr.2113.

3 Pollard D, et al. Standardized Economic Cost Modeling for Next-Generation MAb Production. BioProcess Int. September 2016.

Renaud Jacquemart, PhD, is principal scientist and director of vaccines process sciences; and James G. Stout, PhD, is vice president of process sciences at Natrix Separations, Inc., 5295 John Lucas Drive, Unit 6, Burlington, ON, L7L 6A8 Canada; 1-905-319-2682, fax 1-905-319-0430.

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Depth filtration at Rentschler (WWW.RENTSCHLER.DE)

For current process development phases, many biomanufacturers’ attention is directed increasingly to the first unit operation in downstream processing, which is the removal of cells and cell debris from culture broth and clarification of supernatant containing a biopharmaceutical product. Given the high cell densities achievable with both mammalian and microbial cell culture processes, primary recovery can be a significant challenge.

The current trend in cell culture is to increase product titers with enriched culture media, improved cell productivity, and increased cell mass. High titers also can be achieved through increased culture duration, which can lead to a significant drop in cell viability. These factors cause the increase of process impurities such as host-cell proteins (HCPs), nucleic acids, lipids, and colloids, as well as generation of a broad particle-size distribution in cell culture fluids. Downstream chromatography requires a fast and reliably produced particle-free supernatant.

Centrifugation
Centrifuges perform a robust clarification process. This is a common technique used to harvest large-scale cell culture vessels because it combines low running costs with uncomplicated process development and operating robustness (2–4). Multichamber, tubular bowl, and disc-stack designs all are available. Centrifuges remove a considerable proportion of cells and cell debris that can foul downstream filters and chromatographic steps, leading to unacceptable pressure drops and reduction in overall performance. An optimized centrifugation process minimizes cell lysis — and related generation of additional cell debris or release of intracellular impurities and proteases — and maximizes sedimentation of submicron particles and product yield (5).

Most current harvest operations make use of hermetically sealed, bottom-fed centrifuges that completely eliminate the air–liquid interface commonly present in standard nonhermetic centrifuges. That interface can be harmful to the integrity of the mammalian cells, and it creates additional cell debris when not managed appropriately through backpressure. Fully hermetic centrifuges use mechanical seals that isolate product fluid from outside air, eliminating cell-damaging air–liquid interfaces by enabling the machine to be completely filled with fluid. Such a design also eliminates valves and pumps from the inlet piping, which contributes to an overall reduction in cell lysis by >50% (6, 7).

Single-use options include CARR Centritech’s UniFuge system from PneumaticScaleAngelus. Its gamma irradiated, disposable module eliminates the need for cleaning in place (CIP) and steaming in place (SIP). The unit provides relatively low shear forces that will not compromise the integrity of animal cells. However, a limited feed flow rate of ≤6 L/min constrains this centrifuge to batch volumes of 2,000 L or less, with cell culture harvesting periods of six hours at minimum.

Alternatively, kSep single-use centrifuges from Sartorius Stedim Biotech can handle up to 6,000-L culture volumes with a throughput of 12 L/min. Both systems can be applied for continuous processing, retaining cells in a bioreactor while harvesting culture supernatant. Scale-down models for centrifugation are not always representative, however, which complicates linear scale-up in process development. In addition, centrifugation cannot completely remove the submicron-particle load from a product stream. Depth filtration remains a mandatory step before loading the first chromatography step downstream.

Tangential-Flow Microfiltration
Flow filtration is the simplest procedure for clarifying cell culture supernatants of nearly unlimited scale. Tangential-flow filtration (TFF) is often used based on its ability to counteract continually the formation of a surface layer (preventing associated membrane blockage). A cell suspension is pumped through membrane-bounded channels, and flow is adjusted to operate under quasilaminar conditions.

Most companies use TFF cassette systems with open-channel configuration rather than turbulence-promoting installations or hollow-fiber modules. However, significant shear forces are generated along a TFF membrane to prevent filter blockage. Inevitably those are combined with pressure drops in the flow channel, causing transmembrane pressure variation along the channel length and thus a nonhomogeneous flow-through. Filtrate flow and convective transport of particles to the membrane are significantly higher in regions of high transmembrane pressure than in regions of a low transmembrane pressure. This leads to formation of a surface layer and fouling effects, which increase along the length of the membrane channels during filtration. Thus, the available membrane area is reduced continuously, with increasing time of operation. Shear forces cannot be increased further without negative effects on cell integrity.

Single-use TFF options for application in the biopharmaceutical industry include the Allegro CS system from Pall Life Sciences. It can support cell culture supernatants of ≤2,000-L batch volumes with a maximum cassette surface area of 10 m². Companies maximizing cell culture titers through combined increases in cell density, specific productivity, and culture duration have seen increases in impurity levels and the percentage of solids present in their cell culture fluids. Such increases in submicron debris load made implementation of TFF more challenging, leading some companies to choose centrifugation using heat-sterilizable stainless steel centrifuges coupled with depth filtration. That is the current industry standard for harvesting cell culture fluids from larger — ranging about 5,000–10,000 L — bioreactor volumes.

One TFF variant is Alternating Tangential Flow (ATF) technology from Repligen, which is used primarily for cell retention in perfusion processes. It prevents fouling during long-term applications by creating an alternating flow through the action of a diaphragm, which prevents clogging of the system’s hollow fibers. Recently a presterilized version became available: the XCell ATF6 system, which handles volumes ≤125 L.

Depth Filtration
Cell culture supernatants can be clarified comparatively easily by common-flow (“normal-flow”) filtration. This uses either a membrane with a defined pore size (for dead-end filtration) or a porous material with decreasing pore size along a filter’s thickness (for depth filtration). By contrast with dead-end filtration, in which material is retained on the filter surface, depth filters do not form a filter cake. Additionally, depth filtration can retain particles that are smaller than its pores through a pore-size gradient that separates a broad range of particle sizes (8). Filtration rates differ and are defined by depth-filter manufacturers because no validated standard test method is available. The test for sterile filters, for example, measures their retention of 10⁷ colony forming units (CFU) per milliliter (mL) of the test bacterium Brevundimonas diminuta. When all pores of a depth filter are occupied, a depth filtration process transitions to a dead-end filtration operation.

Depth filters come in several different forms. A common design consists of a layer of cellulose, a porous filter aid such as diatomaceous earth (DE), and a positively charged polymeric resin that binds the two together. Based on those major components, depth filters remove impurities and particulate material through size exclusion as well as adsorption and provide essential protection for membrane filters downstream. For laboratory, pilot, and small-scale clinical applications, users typically place two types of depth filters (a coarse one followed by a tighter one) in series and load them directly with whole cell broth with no centrifugation or TFF step upstream. For larger-scale clinical and commercial operations (>2,000 L), depth filtration is usually coupled with centrifugation.

Depth filters rely on the high porosity of diatomaceous earth (DE). Particles are withheld inside through both sieving and adsorption effects. Typically 10–20 µm in diameter, animal cells are retained mainly through sieving effects; cell debris and small particles need both sieving and adsorption effects for efficient filtration. The adsorption effect also is responsible for binding deoxyribonucleic acid (DNA) and HCPs, but it can bind a protein of interest as well (9).

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Table 1: Overview of process-scale depth filtration systems (according to manufacturers’ specifications); DE = diatomaceous earth

Several depth filtration systems are on the market (10). All process-scale models — e.g., Millistak+ Pod from Millipore Sigma, Stax from Pall Corporation, 3M Zeta Plus from Cuno Inc., and Sartoclear P from Sartorius Stedim Biotech — can separate cells and prepare culture fluid for downstream chromatographies (Table 1). Equivalent small-scale filters also are available for process development and down-scaled models. Area sizes range from 23 cm² to 3.7 m², allowing for free and suitable composition and adaptation to specific volumes and requirements. Process engineers extrapolate the optimal filter area at laboratory scale to a corresponding large-scale filter size. Beyond certain high volumes and particle contents, suitability of depth filtration is limited, but that can be overcome (for example) by sedimentation of culture broth before filtration.

Most depth filters vary by manufacturer in their membrane composition, which is basically cellulose fibers comprising several thousand glucose units that create a fiber labyrinth for retaining particles. Moreover, all manufacturers use filter layers containing DE, which consists of silicon dioxide from the shells of ancient diatoms. DE is characterized by a very porous structure and acid resistance for a high filtration performance. Membranes in 3M Zeta Plus filters additionally contain perlite, which is a volcanic amorphous glass made from crystalline silica (obsidian). Stax filters are made of resins, and Sartoclear P filters contain crosslinking material as a further component.

Process filter cartridges are inserted into a pilot- or a process-scale holder that accepts up to 10 filters per rack. Up to three racks can be used simultaneously for adaptation to any desired scale. These large process filters are not presterilized and must be assembled according to specific process requirements. Filter capsules can be sterilized at 121 °C with 1 bar overpressure for 30 minutes (3M Zeta Plus) or an hour (Stax, Millistak+, Sartoclear P), and they are resistant to sanitization with 1 mol/L NaOH for an hour. Maximum overpressure for operation ranges 1.0–3.5 bar.

Commercially available depth filters can be adapted to any production volume. Encapsulated depth filters from the 3M Zeta Plus EXP SP series have different pore sizes and filter layers. Some units (e.g., 10SP02A, 30SP02A) are suitable for cell separation and protecting chromatographic columns from particles; other filters (e.g., 60ZA05A, 90ZA05A) contain a strong positively charged crosslinking polymer suitable for separating DNA and HCPs. The largest process depth filtration holder allows for a maximum effective filter area (EFA) of 11.2 m² for a single layer (SL) and 17.5 m² for a double layer (DL) to handle cell culture volumes ≤5,000 L. Downstream process engineers can use a multiround holder with several racks in a carousel to clarify nearly any production volume efficiently.

Stax filters differ in the content of two different resins. Wet-strength resins increase membrane stability for higher compressive strength. Charged resins induce a zeta potential within the filter, depending on the culture medium used, which promotes separation of DNA and HCPs. These filters come in single- and double-layer forms. The first (coarser) layer is identical in all DL filters. The second (finer) layers can have different pore sizes. The largest Stax SL filter (EFA 2.0 m²) can clarify cell culture volumes of ≤20,000 L.

Millistak+ Pod filters incorporate multiple graded-density layers and adsorptive, positively charged filter media. The D0HC DL model (EFA 1.4 m²) is used for cell separation in largescale processes. Single-use process filters in the Sartoclear P series come as DL filters for clarifying cell culture supernatants. Large-scale process filter systems of all manufacturers are stackable for use in stainless steel housings.

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Figure 1: Depth filtration was performed on a MAb-producing recombinant CHO cell (20-µm diameter) suspension containing fed-batch culture supernatant from a 10-L Xcellerex XDR-10 single-use bioreactor (GE Healthcare) and a 10-L BIOSTAT B glass bioreactor (Sartorius Stedim Biotech), each with 7.5-L working volume. Final cell concentration was 1.16 × 10⁷ mL^–1, and cell viability was 78%. Total volumes of 4,167 (1) and 4,186 L (2), respectively, were harvested for depth filtration using a 270-cm² Clarisolve 20MS-Millistak POD polypropylene and cellulose/DE filter, a X0HC 270-cm² Millistak POD polypropylene filter, and a 3 × 20 cm² AcroPak 20 cartridge with a Supor EKV membrane made of hydrophilic polyethersulfone (Pall Corporation) with a pore size of 0.2 µm.

Small-scale depth filtration reveals an upscalable down-scaled model. Large-scale depth filtration can be evaluated reliably through laboratory-scale models based on an ensemble of corresponding small filter systems from the same suppliers. Figure 1 shows a depth filtration using a 270-cm² Millistak POD filter series to clarify a 5-L cell culture within 3.5 hours, with results normalized to 1 m² EFA. By achieving the maximum predefined overpressure of 1 bar without filter blocking, the downscale model shows optimal filtration areas for an economical process. Based on these results, we calculate a minimum filter area of 6.2 m² and 5.1 m² for 1,000-L cell culture for a Clarisolve 20MS-Millistak POD filter and a X0HC Millistak POD membrane, respectively.

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Figure 2: Depth filtration was performed on a MAb-producing recombinant CHO cell (20-µm diameter) suspension containing fed-batch culture supernatant from a 2,000-L Xcellerex XDR-2000 single-use bioreactor (GE Healthcare) with a final cell concentration of 1.38 × 10⁷ mL^–1 and 74% viability. Three consecutive depth filtration runs used a 3M Zeta Plus encapsulated three-filter system with a respective 16EZB large filter holder consisting of a 1.6-m² cellulose/DE filter with a pore size range of 1.5–10.0 µm, a second 1.6-m² cellulose/DE filter with a pore size range of 0.2–2.0 µm, and a 1.56-m² hydrophilic PES microfilter with a pore size of 0.2 µm. Clarified culture supernatants were harvested in 1,000-L Mobius single-use process containers made of PureFlex film (Millipore Sigma).

Large-scale depth filtration systems operate with high filtration performance throughout an entire harvesting process. For routinely increasing operational safety, cells are sedimented for 12–22 hours at 10 °C before depth filtration. Figure 2 shows the final three representative processes of a depth filtration for 2,000-L volume from a high–cell-density culture supernatant of a fed-batch cultivation. Every filtration run harvested filtrate of about 500 L of culture supernatant. The graph shows an identical slope for each run at a constant flow rate of 12–13 L/min. For the first filtration (into Filtrate Container 1) the flow rate increased gradually from 5 L/min to 12 L/min within the first 15 minutes of operation. No reduction in performance was observable during the whole clarification process using EFAs of 1.6 m² and 1.56 m², with 2,000 L harvested in about three hours.

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Table 2: Comparing single-use harvest clarification techniques

A Bright Future
A number of single-use technologies for harvest clarification — centrifugation, TFF, and depth filtration — have different advantages and disadvantages (Table 2). We find depth filtration to be highly preferable because of its low investment cost, good scalability, easy handling, and reproducible clarification efficiency. Process improvement is ongoing through optimization of primary recovery without time-consuming sedimentation steps. For example, harvest material may be pretreated with flocculants (such as polydiallyldimethylammoniumchloride) or acidification (11, 12). Meanwhile, suppliers are developing depth filters for special flow-through, further strengthening the value of this single-use harvesting technique.

References
1 Singh V. Disposable Bioreactor for Cell Culture Using Wave Induced Agitation. Cytotechnol. 30, 1999: 149–158.

2 Axelsson H. Cell Separation, Centrifugation. Encyclopaedia of Bioprocess Technology. Flickinger M, Drew S, Eds. Wiley VCH: Weinheim, Germany, 1999: 513–531.

3 Abraham S, et al. Strategies for Improving Mammalian Cell Clarification. Abstr. Pap. Am. Chem. Soc. 225, 2003: BIOT-119.

4 Hanle D. Centrifuges, Animal Cells. Encyclopaedia of Bioprocess Technology. Flickinger M, Drew S, Eds. Wiley VCH: Weinheim, Germany, 1999: 553–559.

5 Kempken R, Preissmann A, Berthold W. Assessment of a Disc Stack Centrifuge for Use in Mammalian Cell Separation. Biotechnol. Bioeng. 46, 1995: 132–138.

6 Schmidt M. Antibody Degradation (Disulfide Reduction) in CHO Production Process. Antibody Development and Production conference. IBC Life Sciences: Westborough, MA, 2009.

7 Liu HF, et al. Recovery and Purification Process Development for Monoclonal Antibody Production. MAbs 2, 2010: 480–499.

8 Fiore J, Olson WP, Holst S. Depth Filtration. Methods of Plasma Protein Fractionation. Curling J, Ed. Academic Press: London, UK; New York, NY; 1980: 239–268.

9 Yigzaw Y, et al. Exploitation of the Adsorptive Properties of Depth Filters for Host Cell Protein Removal During Monoclonal Antibody Purification. Biotechnol. Prog. 22, 2006: 288–296.

10 Shukla A, Gottschalk U. Single-Use Disposable Technologies for Biopharmaceutical Manufacturing. Trends Biotechnol. 31, 2013: 147–154.

11 Brodsky Y, et al. Caprylic Acid Precipitation Method for Impurity Reduction: An Alternative to Conventional Chromatography for Monoclonal Antibody Purification. Biotechnol. Bioeng. 109, 2012: 2589–2598.

12 McNerney T, et al. PDADMAC Flocculation of Chinese Hamster Ovary Cells: Enabling a Centrifuge-Less Harvest Process for Monoclonal Antibodies. MAbs 7, 2015: 413–427.

Stefan R. Schmidt is vice president of process science and production, Stefan Wieschalka is senior scientist in process science, and Roland Wagner is head of production at Rentschler Biotechnologie GmbH, Erwin-Rentschler-Str. 21, 88471 Laupheim, Germany.

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Single-step harvesting of a high-density cell culture from a Biostat STR bioreactor with Sartoclear Dynamics body-feed filtration from Sartorius Stedim Biotech (WWW.SARTORIUS.COM)

Downstream processing has been considered a “bottleneck” in the manufacture of protein biotherapeutics ever since cell culture engineers began dramatically improving production efficiencies around the turn of the century. And as single-use technologies have grown in importance and acceptance, offering more solutions every year, their biggest challenges too have been in the separation, purification, and processing that follows product expression in cell culture. Many of the technologies familiar to process engineers — e.g., centrifugation and chromatography — present technical and economic problems.

In a recent white paper (1), BioPlan Associates reported that respondents to its 13th annual survey of biomanufacturers reported downstream processing as requiring the most technological improvement of any part of bioprocessing. But productivity is improving. For the coming year, less than half (45%) of respondents expect capacity problems — compared with 88% of the industry in 2005.

Where would user companies like to see suppliers direct their development efforts? Nearly three quarters of respondents (especially contract manufacturers) cited downstream continuous bioprocessing and disposables for purification. Our featured report in May will address the former more specifically — but in general, although interest is high, not many relevant products are available yet. In 2016, bioprocessing facility budgets were up 5% for downstream technologies. “Essentially, nearly all respondents want improvements in downstream technologies,” the report claims (1).

Separation and purification technologies are slowly catching up to upstream processing, however, and vendors are filling the gaps in their offerings. Filtration continues to advance, of course, with some options even encroaching on the adsorption mechanics of chromatography. And now even drug-product filling operations can choose single-use options.

Harvest and Clarification
Modern bioproduction technologies have given us expression titers measured in grams per liter of culture — whereas before the turn of the century, milligrams per liter were common. High-density culturing is one reason, but other advancements include more complicated media and feeds, culture strategies, and optimization efforts. Some of those improvements upstream can cause trouble for harvest, clarification, and beyond.

The first phase of downstream processing typically includes centrifugation or primary filtration steps followed by secondary filtration before purification involving chromatography (2).

Harvesting here is just the same as in agriculture: collecting the material produced by your hard-working life forms (in this case, animal cells or microbes). Along with the expressed protein of interest come host-cell proteins that are interesting only as contaminants; nucleic acids; leftover nutrients, supplements, and byproducts; secondary metabolites; and water. Clarification follows to prepare this messy product stream for downstream chromatography and purification. With solids making up 3–5% of the culture volume, the clarification process alone typically takes the yield of active protein down by 10–15% (2). The most demanding processes involve high turbidities and increased particle/contaminant loads as well as high densities and titers (3).

Filtration: Complications related to downstream processing are ironic in one sense: Disposability in bioprocessing pretty much began with filter cartridges, although upstream/production single-use applications surpassed downstream technologies along the way. Filters remain the most popular single-use technologies, with 91% of companies currently using disposables citing filter cartridges in the latest BioPlan Associates survey, with robust growth continuing (1). Use of depth filters is are widespread (82%), although only 5% of respondents report using disposable tangential flow filtration (TFF) devices.

Another up-and-coming filtration technology is “body-feed filtration,” which incorporates filter-aids such as diatomaceous earth (DE) to increase filter capacity — of particular use with the highly concentrated feed streams that can come from high–cell-density cultures. Adsorptive depth filtration (ADF) incorporates DE into the filter itself (3). Depth filters have particular utility in clarification, which increases with the help of filter aids, flocculating agents that settle impurities out of a harvest solution, or protective prefilters. Sartorius Stedim and MilliporeSigma are well-known proponents of such approaches (2–4). See the next article in this featured report for more discussion.

Filters play many roles in downstream processing — beyond harvest and clarification to virus reduction, buffer exchange, volume reduction, and final sterile filtration just to name a few. Filtration systems are highly automatable, as well (see the box, below), which is increasingly needed in modern biomanufacturing.

Centrifugation has been problematic for conversion to single use, but some suppliers do offer solutions. Most notable are kSep systems (now part of Sartorius Stedim Biotech) and Carr Centritech UniFuge systems from PneumaticScaleAngelus. The former are based on fluidized-bed centrifuge technology originally developed by KBI Biopharma; the latter comprise more traditional technology with irradiated and disposable product-contact surfaces.

The Sartorius technology can be used both for harvesting cells as product or discarding them as by-products. Balanced centrifugal and fluid-flow forces retains particles as a concentrated fluidized bed under a continuous flow of media or buffer. Some companies are applying it toward continuous processing. The Carr system offers continuous operation as well. Both are highly automatable, although they face limitations in scalability and process monitoring (5).

Other Options: Harvest clarification methods such as feed pretreatment involve single-use technology only in that they require tubing to move harvested material and treatments to and from a mixing system (which may or may not be disposable). Acids and salts can cause solutes to precipitate out, but they also can denature proteins; cationic polymers bind contaminants together into cloudy flocs that can be filtered out, but the polymers themselves become contaminants that must be removed later. If such methods are used, they are likely to be combined with the above technologies, whether single-use or multiuse forms thereof.

Chromatography
When you ask about single-use chromatography, the answer usually comes in the form of prepacked columns (e.g., ReadyToProcess brand from GE Healthcare, OPUS columns from Repligen, and Chromabolt and Mobius FlexReady brands from MilliporeSigma). They aren’t strictly single-use in nature, though: Most resins, gels, and other chemistry-based separation media are too expensive to use only once. Instead, they are washed and equilibrated for repeated use with the same product stream, then discarded along with their polymer columns once a batch is complete. In addition to the usual benefits related to cleaning and cleaning validation, however, this approach saves users the time, cost, and fussiness of column packing — giving them consistent results from expert suppliers instead.

Column volumes currently available (e.g., ≤20 L) require several cycles to purify a 1,000-L batch. And that takes time, thus adding cost. The alternative of adding more columns also costs more — lots more — unless you’re talking about continuous multicolumn processes such as that described in this special section by authors from CMC Biologics and Pall Life Sciences. The key with disposable chromatography is to balance cost of goods (CoG) of materials and time/labor. The more expensive the medium (e.g., protein A affinity resins), the more sense it makes to use traditional multiuse technology. So too with frequent harvesting and media with long lifetimes. However, mixed-mode sorbents and sequential chromatography are improving performance while reducing costs. Meanwhile, smaller columns are becoming more popular thanks to high-capacity, high-flow resins and smaller production batches (e.g., from high-density cultures).

A new column-free chromatography technology is drawing publicly stated interest from companies such as Medimmune (Astrazeneca) and Regeneron: Continuous Countercurrent Tangential Chromatography (CCTC) from ChromaTan Corporation. A slurry of resin sequentially binds product as it flows through a series of mixers and hollow-fiber membranes, where it is also washed, eluted, and stripped in a continuous process. The “countercurrent” refers to buffers flowing through in the opposite direction, both lessening the amount of buffer used and improving resin use efficiency. Like the Pall process highlighted elsewhere in this report, CCTC has potential for continuous processing — and more on this small company’s progress is coming in our May featured report.

Alternatives to chromatography resins — in columns or otherwise — are available from membrane suppliers. Functional filtration and membrane adsorbers typically are limited in dynamic binding capacity (DBC) compared with column/resin technology, but they handle significantly higher flow rates.

Nearly one in five respondents to BioPlan Associates’ 13th annual survey cited chromatography columns as currently causing them significant or severe capacity constraints (1). Membrane adsorbers, however, have yet to take over a significant portion of the market. But their adoption is growing, with first use in respondents’ facilities reportedly up 13% for 2016 and 10-year market growth of 31%. The concept is not entirely new technology, but recent introductions — e.g., salt-tolerant devices and new ligand technologies — offer improvements in robustness and efficiency. The article from Renaud Jacquemart and James G. Stout in this special insert provides more discussion.

Formulation, Fill and Finish 
Mixing and storage systems are another single-use technology making inroads with biomanufacturers, at least in part because of their many uses. Their annual adoption rate was up 16% for 2016, and their 10-year growth has been about 50% (1). Downstream applications include viral safety (e.g., detergent treatment), storage of process intermediates, and product formulation, to name a few.

Fill and Finish: BioPlan Associates identified single-use technology as the key trend in biopharmaceutical fill and finish operations, with nearly two-thirds of their respondents ranking it number one (1). Over a third of respondents plan to implement new fill–finish technologies at their facilities in the next two years.

Major suppliers such as MilliporeSigma and Pall have introduced solutions to meet this need, and companies such as Biotest, Disposable-Lab, and Merck have implemented them (6–8). The basic idea has been to adapt fluid-path technology with metal filling needles, combining them with bag containment and pumping systems. As you’ll see in the interview at the end of this featured report, facility design and engineering firm NNE Pharmaplan is a major proponent of these ideas.

Pumps are important throughout downstream bioprocessing, of course. And although polymer tubing and connectors are established single-use components for fluid handling, pumps thus far have proven to be more of a challenge (9). Solutions are in the works to address these needs, and companies such as PSG Dover already have put forth some options. It offers a line of positive displacement diaphragm pumps (Quattroflow) with product-wetted plastic chambers that can be replaced. Other options include a rotary pump from Quantex Arc and a disposable pump-head system for Masterflex peristaltic pumps from Cole Parmer.

Challenges Yet to Be Overcome
Finally, another gap that remains to be filled completely for downstream processing relates to sensing and sampling. Most such solutions that have been made available so far are meant for upstream production applications. PendoTech offers pressure sensors, however, as well as those for ultraviolet absorbance and conductivity (10). Sartorius Stedim Biotech has incorporated such options into its own product offerings (11).

But what might be the biggest challenge facing downstream processors who want to use disposable systems and components isn’t so much technical as it is business related. What users really want from their single-use technology providers is standardization of designs that would allow them to mix and match components to put together systems that work best for their own processes. Bioprocessors say that this would improve adoption and implementation of disposables; suppliers are reluctant to share with their competitors. Not long ago, in fact, if you brought up the question of single-use standardization at an industry conference, you might get a lot of laughs but not much real discussion. However, companies on both sides are taking the idea more seriously now. And the “alphabet soup” of organizations concerned with single-use technology are helping to make it happen (12). Standards could reduce the risk of process failures and allow suppliers to stick with proven features while focusing their attention on needed innovations.

References
1 Top 15 Trends in Biopharmaceutical Manufacturing, 2016. BioPlan Associates, Inc.: Rockville, MD, 2016.

2 LeMerdy S. Evolving Clarification Strategies to Meet New Challenges. BioProcess Int. October 2014: insert.

3 Minow B, et al. High–Cell-Density Clarification By Single-Use Diatomaceous Earth Filtration. BioProcess Int. 12(4) 2014: S36–S46.

4 Schreffler J, et al. Characterization of Postcapture Impurity Removal Across an Adsorptive Depth Filter. BioProcess Int. 13(3) 2015: 36–45.

5 Pattasseril J, et al. Downstream Technology Landscape for Large-Scale Therapeutic Cell Processing. BioProcess Int. 11(3) 2013: S38–S47.

6 Camposano D, Mills A, Piton C A Single-Use, Clinical-Scale Filling System: From Design to Delivery. BioProcess Int. 14(6) 2016: 50–59.

7 Gross R, et al. Establishing Single-Use Assemblies on Filling Equipment. BioProcess Int. 12(4) 2014: S48–S54.

8 Zambaux J-P, Barry J. Development of a Single-Use Filling Needle. BioProcess Int. 12(5) 2014: 46–53.

9 Wittkoff W, Prasad R. Single-Use Pumps Take Center Stage. BioProcess Int. 11(4) 2013: S18–S23.

10 Annarelli D. Novel Single-Use Sensors for Biopharmaceutical Applications. BioProcess Int. 12(7) 2014: 58–59.

11 Weichert H, et al. Integrated Optical Single-Use Sensors: Moving Toward a True Single-Use Factory for Biologics and Vaccine Production. BioProcess Int. 12(8) 2014: S20–S24. 1

2 Vogel JD, Eustis M. The Single-Use Watering Hole: Where Innovation Needs Collaboration, Harmonization, and Standardization. BioProcess Int. 13(1) 2015: insert.

Further Reading
Bird P, Hutchinson N. Automation of a Single-Use Final Bulk Filtration Step: Enhancing Operational Flexibility and Facilitating Compliant, Right-First-Time Manufacturing. BioProcess Int. 13(3) 2015: S40–S43, S52.

Blomberg M. The New Hybrid: Single-Use Systems Enabled by Process Automation. BioProcess Int. 13(3) 2015: S34–S39.

Grier S, Yakabu S. Prepacked Chromatography Columns: Evaluation for Use in Pilot and Large-Scale Bioprocessing. BioProcess Int. 14(4) 2016: 48–53.

McGlaughlin MS. An Emerging Answer to the Downstream Bottleneck. BioProcess Int. 10(5) 2012: S58–S61.

Metzger M, et al. Evaluating Adsorptive Filtration As a Unit Operation for Virus Removal. BioProcess Int. 13(2) 2015: 36–44.

Mok Y, et al. Best Practices for Critical Sterile Filter Operation: A Case Study. BioProcess Int. 14(5) 2016: 28–33.

O’Brien TP, et al. Large-Scale, Single-Use Depth Filtration Systems. BioProcess Int. 10(5) 2012: S50–S57.

Quinlan A. Advances in Chromatography Automation. BioProcess Int. 13(1) 2015: 16–17.

Cheryl Scott is cofounder and senior technical editor of BioProcess International, PO Box 70, Dexter, OR 97431; 1-646-957-8879; cscott@bioprocessintl.com.

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Figure 1: Harvest clarification options

Steadily increasing demand for biopharmaceutical drugs has led the industry to examine its manufacturing scales while pressuring research and development groups to produce high-yielding clones and processes. Improved media, feed supplements, bioreactor designs, and control of process parameters have helped biomanufacturers achieve multifold increases in volumetric productivity from production bioreactors.

However, cell culture processes are significantly affected by their bioreactor’s ability to support cells at higher densities and sustain cultures at lower viabilities. With the implementation of a number of new approaches, cell densities have been increased from 5–7 × 10⁶ cells/mL to >25 × 10⁶ cells/mL. Such increased densities — and improved specific productivity of the modern clones — has increased productivity as well, with expression titers rising from 1.0 g/L to 5–8 g/L in fed-batch cell culture processes. Moreover, concentrated fed-batch and perfusion cell culture processes have further increased cell density up to >50 × 10⁶ cells/mL, which has further complicated clarification as well as overall downstream processing of recombinant proteins. A high-density cell culture process always poses a challenge to clarification techniques for separating product from cells without compromising its yield or quality.

In addition to cell culture processes, clarification and purification techniques are under tremendous pressure to deliver high-quality biological products while controlling cost of goods (CoG). Choice of clarification technique directly affects final-product quality because it paves the way to downstream purification processes that build product quality and ensure product safety.

The basic intention of a cell-separation unit operation is to remove whole cells and their components for further processing of a culture’s supernatant. No single technology fits all needs of such applications for all products and processes. The choice of clarification technique varies for different cell densities and viabilities, cell type, culture styles and modes, and harvest viscosities. As cell density and product titer have improved over the past decade, clarification techniques also have improved to meet resulting process and product requirements. We recommend screening all commercially available technologies to understand the best available option for manufacturing of a given biopharmaceutical. We also prefer single-use devices, performing screening studies of high-capacity depth filters as reported herein.

The objective of the study described below was to evaluate different single-use technologies and understand the impact of high cell density and lowered viability on existing devices. We challenged different available depth filters with similar feed streams to evaluate and compare their performance.

Impact of High Cell Density and Low Viability
The harvested supernatants of today’s fed-batch processes, with very high cell densities and low viabilities, show increased solid materials such as whole cells, cell debris of different sizes, colloids (lipid components of media and cell walls), nucleic acids, and other particulate matter. Those increased levels can cause early plugging of traditional depth filters. In many cases, multifold increases in filter area are required, leading to poor scalability and expense at manufacturing scale.

Traditional primary clarification techniques such as centrifugation, tangential-flow filtration (TFF), and early single-use depth filtration have been used in biomanufacturing to achieve desired quality outputs. They are still integral to many commercial manufacturing processes. However, downstream process engineers are looking for alternatives that can provide more efficient clarification of high-density cultures. Combinations of the familiar techniques have been applied in some cases, depending on the need of the cell culture process and its scale of operations (1). All these techniques — or combinations thereof — have served us at some point. Figure 1 illustrates some combinations of clarification options.

Most downstream processes in biopharmaceutical manufacturing include two major categories of technology: chromatography and filtration. The latter includes clarification, virus filtration, and tangential-flow filtration. Clarification process can be separated further into two broad categories: primary recovery and secondary recovery, also referred as primary and secondary clarification steps. The primary step removes the bulk of large particles, whole cells, and cell debris; the secondary step removes smaller particles present in the resulting filtrate. Centrifugation, TFF, and depth filtrations are common choices for primary clarification; depth filtration and bioburden-reduction filters often provide secondary clarification.

Primary Clarification Options
Tangential-flow microfiltration (TFF-MF) separates particles based on size exclusion using microfiltration membranes with a pore size of ≤0.65 μm. This process is highly efficient for removing whole cell mass and fragments. It provides the most consistent separation by retaining particle sizes larger than the membrane cutoff size.

TFF-MF devices also offer advantages for scaling up downstream processing based on modularity. With higher cell densities, however, ruptured and fragmented cells often are observed in recirculation loops, making a secondary clarification necessary to reduce those smaller particles before further downstream processes (e.g., chromatography). High cell density can increase polarization on cell membrane surfaces, leading to frequent filter clogging and complicating manufacturing-scale operations.

High-yielding cell culture processes with higher cell densities and specific productivities produce large amounts of solid material (>6%). TFF is best suited for such cultures with solid content <3%. Beyond that, however, TFF becomes inefficient.

Centrifugation can be applied to feed streams with high solids levels, concentrating cell mass ≥40% v/v. Product recovery can be low, however, because of increased pellet volumes and high desludging, which are common for cell harvests with very high solids content. Additionally, low cell viability generates particles of different sizes, drastically decreasing their removal efficiency. Although this technique can handle high concentrations of insoluble materials, its ability to produce a clear product is limited. Thus, it often requires the support of secondary filters.

In most cases, loading a cleaner material on a column requires depth filtration after centrifugation and before chromatography. The area requirement for such a depth filter would depend on the clarity of the centrifugation output and the amount of suspended material present after primary clarification.

Depth filters typically are made of cellulose, with a porous filter aid such as diatomaceous earth (DE) and an ionic charged resin binder. These filters function by retaining particles within their porous matrices. Depth filtration has been a single-use choice for clarification in manufacturing many biopharmaceutical products.

Depth-filter pore sizes vary for primary and secondary applications. Different types of depth filters are now available with single or multiple layers and gradient pore distributions. These filters can be applied either directly to whole cell broth (for cleaner output) or used in two stages: primary depth filters to remove larger particles, followed by secondary depth filters to remove finer suspended particles. Depth filters fit well with the emerging trend of single-use technology, making them our primary choice as a single-use clarification solution in biopharmaceutical development.

With the advancement of single-use technology, primary and secondary filters are merged into one single step that reduces cycle times and required filter area for efficient clarification. Furthermore, lower buffer volumes are required for flushing in modern processes, enabling improved economics for clarification of high-density cell cultures. Finally, hardware requirements for depth filtration at higher scale are simple and highly flexible, which makes depth filters an attractive choice as a single-use clarification technique.

Key features needed for a filter to qualify its use from laboratory scale through pilot to commercial scale are high capacity with minimal area, low cost, consistent performance, high product quality and yield, scale-up ease and flexibility, and a small overall footprint for efficient use of space in a biomanufacturing facility.

Depth Filter Study
For evaluation of existing depth filters and other improved technologies, we divided our experiments into two phases. In phase 1, we obtained and evaluated available depth filters and additives capable of handling higher cell densities from different vendors. Filters were challenged with ~28 × 10⁶ cells/mL of cell culture harvest having ~75% cell viability. We analyzed the performance of all filters to select a few best candidates for our phase 2 study. Then, we challenged those shortlisted filters with a bit lower cell density (~24 × 10⁶ cells/mL) and even lower viability ~65%. Phase 2 was meant to study reproducibility and robustness of filter performance with some variation in an upstream process.

In most bioprocess applications using depth filters, flow rate in relation to membrane cross-section area has been found to affect process capacity significantly (1). It’s now known that the increased flux reduces the capacity of a membrane (L/m²) (1). Therefore, to achieve acceptable depth-filter performance, we applied a constant flow method. This keeps flux throughout a process, during which pressure and turbidity are monitored until they reach their maximum allowable operating limits. We used different endpoints for each filter, as specified and recommended by its supplier. Finally, we compared data from all experiments and interpreted them to determine the best-suited depth filtration technique for our process.

Materials and Methods
In our phase 1 study, we evaluated depth filters from four suppliers — Pall Life Sciences, Sartorius Stedim Biotech, MilliporeSigma, and 3M — with different pore sizes and multiple layers or both. Each filter was tested for its ability to reduce turbidity to a desired level: <15 NTU, preferably <10 NTU. We used a LaMotte turbidity meter throughout this experiment to measure sample turbidity.

We also compared an alternative clarification technology: flocculation, which relies on aggregation of particles for easier separation. In this method, a cationic polymer is added to harvested supernatant to trigger floccule formation. The polymer binds to whole cells, cell debris, and other components, then attracts other nearby components by the van der Waals force to form aggregates of varied size and density. We tested this method along with depth filters of different pore sizes as recommended by their vendors. The polymer dose varies from harvest to harvest, and it’s important to determine the optimum concentration of flocculant for each cell culture process. To determine an appropriate concentration, we added an increasing volume of flocculant solution to a constant volume of harvest, checking all samples to determine which reached the lowest turbidity. We thus considered the concentration at which the sample showed lowest turbidity to be the optimum flocculant concentration for filtration. Then we challenged a harvest with this optimum concentration further using depth filters of different pore size to discover the best-performing depth filter.

Finally, we evaluated another technology: addition of diatomaceous earth (DE) powder to harvested supernatant before filtration. This technique increases the surface area of a depth filter by making a porous cake of DE over it. In normal depth filtration, a filter cake forms on the surface of a filter and clogs its available surface area. But with DE added, it forms a permeable layer over the filter, which increases its depth and surface area for filtration. The porous layer traps whole cells and debris through both adsorption and sieving to yield a clear filtrate. Again, the amount of DE required varies with harvested cell mass. It can be determined by measuring pelleted cell weight and using the result to calculate a recommended percentage of DE. We also assessed this method’s performance using recommended filters.

So the different types of single-use clarification technologies we evaluated are as follows: a single depth filter (single-stage clarification), primary and secondary depth filters (two-stage clarification), flocculation followed by depth filtration, and DE additive (to enhance filtration efficiency) followed by filtration.

Results and Discussion

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Figure 2: Comparative profile of all the depth filters at constant flow (phase 1 study)

Figure 2 shows a comparative profile of all filters tested in phase 1, plotting the correlation of filter performance (capacity) with differential pressure. Table 1 summarizes the values of reduced turbidity and capacity for those filters. And Figures 3A and 3B provide a comprehensive summary of the phase 1 study.

We performed two different experiments on filters provided by Pall Life Sciences. In the first experiment, the HPPDH4 depth filter showed a capacity of 47.3 L/m² and a pooled turbidity of <12 NTU. The second experiment used primary and secondary depth filters SC050PDK5 and SC050PDE2.

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Table 1: Performance comparison of depth filters (phase 1)

The main aim of the first-stage filter SC050PDK5, with its wider pore size, is to remove larger particles with a turbidity cutoff of <25 NTU. Its output then goes to the secondary depth filter SC050PDE2 (narrow pore size) to get a final pooled turbidity value of <10. Capacity for the primary filter was 63.6 L/m²; a correct capacity of secondary filter SC050PDE2 could not be determined because of limited output from primary filter.

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Figure 3: Comparative performance of depth filters from different vendors (phase 1)

Among the MilliporeSigma filters, those having wider pores (60 HX) showed a capacity of 143 L/m² with no increase in pressure, whereas the 40 MS filter with intermediate pore size showed a very high capacity of 383 L/m², and the 20 MS filter with the smallest pores showed a filtration capacity of 315 L/m², within the allowed pressure limit. Turbidity reduction was acceptable for both the 20 MS and 40 MS units, but the 20 MS filter showed lower pool turbidity than the 40 MS filter.

In our DE-based filtration study, we mixed 40% and 50% DE material from Sartorius Stedim to the harvest cell weight. Results suggest that the harvest with 40% DE showed a better capacity and turbidity reduction than the harvest with 50% DE addition. Final turbidity with both harvests was comparable and >10 NTU.

Within a set pressure limit, we used a primary depth filter (10SP02A) provided by 3M to generate material for a secondary depth filter (90ZB08A). The first showed a low capacity value of 80 L/m² and a high turbidity value of 264 NTU; the second, with its tighter pores, showed a better capacity of 130 L/m² and pooled turbidity of <5 NTU.

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Figure 4: Comparative performance of selected depth filters phase 2

To select the best-performing filters for our phase 2 study, we considered the capacity, turbidity reduction, and process recovery data from all filters used in the phase 1 study. Two new technologies — from Sartorius Stedim and MilliporeSigma — showed promising results for handling high-density cell culture harvests. Therefore, we used Sartoclear Dynamics with 40% DE added and Clarisolve 20 MS and 40 MS filters for our phase 2 study (Figures 4A and 4B).

At the end of the clarification study, we recovered the filter hold-up by allowing air to push product out of the filter to prevent further dilution. Alternately, hold-up volume can be recovered by flushing a filter by two or three hold-up volumes of buffer. Product recoveries for the selected filters were nearly comparable at ≥95%. Actual recovery of the Seitz HPPDH4 primary filter could not be determined, however, due to clogging of primary filters that occurred during material generation.

Overall performance of selected filters in this case was comparable at small scale. But we recommend calculating and optimizing recoveries at pilot scale during scale-up. Pilot-scale batches requires an additional cassette holder, tubings, valves, and connectors — all of which further increases the hold-up volume of filter cassettes.

We found the performance of both Clarisolve 20 MS and 40 MS filters to be comparable and better than that of Sartoclear 40% DE. We observed an increased capacity for all three options than what was seen in our phase 1 results, which could be attributed to a decrease in cell density. But turbidity reduction for all filters in phase 2 was comparable and consistent with our phase 1 data.

An Upcoming Technology
Acoustic wave separation promises to handle very high cell densities from both concentrated fed-batch and perfusion cultures, in which cell density can reach >50 × 10⁶ cells/mL of harvest. We look forward to testing the new Cadence Acoustic Separation (CAS) technology from Pall Corporation as a potential solution to the associated with a very high-density cell cultures.

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Figure 5: (A) Direction of cell movement and increase in size of cells at acoustic zone; (B) how accumulated cells at the base are collected and removed as sludge

In a CAS system, continuous standing waves retard the flow of cells. As cells increase in number, they aggregate into bigger and bigger clumps. Those experience higher gravitation and cannot remain in the flow, so they settle to the conical end of the system’s flow cells (Figure 5). Multiple acoustic flow-cell chambers in series can further reduce turbidity as required for a given process. This seems to be a good option for cell removal in continuous processing as well as harvest of very high-density cell cultures. However, a secondary depth filter may be required to decrease turbidity further downstream of the CAS system.

An Iterative Approach
Several new technologies are available now for clarification and handling high-density cell culture harvests. But no single technique fits all cell culture processes and products. Therefore, it is important to evaluate all available technologies to determine the best clarification solution for an efficient and economical biomanufacturing process. For example, turbidity reduction and filter capacity are important criteria in selection of single-use depth filters for clarification purposes.

In our study, Clarisolve filters (20 MS and 40 MS) showed better performance than others tested. However, the choice of clarification technology for any commercial application must be critically evaluated before its implementation in a final process. Pros and cons of each technology and their implications in a given manufacturing process must be understood at laboratory scale — and preferably at pilot scale — before implementing a final choice at commercial manufacturing scale.

Furthermore, other factors also should be taken into consideration: e.g., cost of each technology, facility requirements, hardware needs, process scalability and flexibility, vendor support, and regulatory requirements. In our study, we found that the higher-density cell culture could be handled with existing depth filtration techniques to get a desired end product. But an extensive study of available options is the most appropriate way to determine your own needs.

Acknowledgments
The authors are thankful to Ipca Laboratories Ltd. (Mumbai, India) for necessary support. They also give special thanks to Dr. Ashok Kumar (president of Ipca laboratories Ltd.) for his immense motivation and necessary support.

References
1 Yavorsky D, et al. The Clarification of Bioreactor Cell Culture for Biopharmaceuticals. Pharm. Technol. March 2003: 62–76.

Further Reading
Collins M, Levison P. Development of High Performance Integrated and Disposable Clarification Solution for Continuous Bioprocessing. BioProcess Int. 14(6) 2016: S30– S33.

Dhanasekharan K, et al. Emerging Technology Trends in Biologics Development: A Contract Development and Manufacturing Perspective. BioProcess Int. 14(9) 2016: 32–37.

Gronemeyer P, Ditz R, Strube J. Trends in Upstream and Downstream Process Development for Antibody Manufacturing. Bioengineering 1(4) 2014: 188–212.

Le Merdy S. Evolving Clarification Strategies to Meet New Challenges. BioProcess Int. 12(9) 2014: I10–I12.

Pegel A, et al. Evaluating Disposable Depth Filtration Platforms for MAb Harvest Clarification. BioProcess Int. 9(9) 2011: 52–54.

Tomic S, et al. Complete Clarification Solution for Processing High Cell Culture Harvests. Sep. Purif. Technol. 141, 2015: 269– 275.

Manish K. Sharma is senior manager, Snehal Raikar is a research associate, Smriti Srivastava is a junior research associate, and corresponding author Sanjeev K. Gupta is general manager of the advanced biotech laboratory at Ipca Laboratories, Ltd. in Mumbai, India; +91-22-6210-5820; sanjeev.gupta@ipca.com; www.ipca.com.

The post Examining Single-Use Harvest Clarification Options: A Case Study Comparing Depth-Filter Turbidities and Recoveries appeared first on BioProcess International.

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Figure 1: Conventional and M-pleat pattern

Maximizing filtration-area density is a design strategy to minimize filter footprint and improve filtration process economics. Pleated membrane formats commonly are used to achieve that goal for sterilizing-grade filters operating in dead-end mode (also known as normal-flow filtration). Although high-density pleat geometries increase productivity for a device, such formats can present unique challenges. One of the most common concerns is that pleat formats can introduce flow resistance that impedes a device’s filtration efficiency, particularly for high–pleat-density geometries (1, 2). Filtration efficiency can be affected by pleat length, membrane permeability, and thickness and flow characteristics of upstream and downstream supports as well as other aspects of a pleat configuration.

One approach for overcoming those challenges has been the development of an M-pleat pattern that allows for a nearly 100% increase in membrane area over conventional designs (Figure 1). The M-pleat design has been optimized for maximum area density, device manufacturability, and a high level of device robustness. Other improvements include the ratio of long to short pleats, pleat compression, and upstream and downstream support materials (see “Design Improvements” box below). M-pleat designs help address the challenges of filtration efficiency while matching existing cartridge sleeve and length dimensions.

Ultimately, filter-design efficiency depends on how well a membrane performs when it is in a filtration device. One method of measuring a device’s filtration efficiency is to compare the performance of a membrane in a filtration device with that of one in a flat-sheet format (where the flow resistance is dominated by the membrane itself). The ratio of commercial-scale filter performance to small-scale, flat-sheet performance is often referred to as the scaling factor. Ideally, that should be at or close to one.

The scalability factor of a device must be defined with respect to a set of specified conditions. In particular, there can be a compromise among other filter requirements (including high-area density) and good scalability. Scalability also can be sensitive to filter operating conditions, challenge stream characteristics, and the particulars of a selected filtration endpoint. A filter can exhibit a range of scaling factors depending on the combination of those other conditions.

Here we discuss variations in scalability in newly developed high-area versions of sterilizing-grade filters. We describe a model that was developed to predict filter device efficiency as a function of the filtration properties of a filtered stream. The model can be used to assess which applications can benefit most from high-area pleated devices and which should use conventional pleat configurations.

Our study evaluated filters for scalability against a nonplugging stream (water) and three different plugging streams that represented a wide range of particle-size distributions. We assessed scalability as a function of particle size and degree of plugging. Filters were tested primarily for constant pressure operation and measured for constant-flux operation.

Theoretical Background
Several factors must be considered when scaling from discs to pleated devices. Applicable to all pleated filter devices, these factors include flow resistance from upstream and downstream supports, pressure losses from housings and plumbing, and variability in filter properties, stream characteristics, and operating conditions (3). Although some theoretical treatments of flow restrictions in conventional pleat patterns have been studied, no known similar treatments have been published for the M-pleat pattern evaluated in this work. Here we focus on the effect of the M-pleat pattern on filter performance, because factors such as membrane and process variability are not specific to high-area devices.

Rather than develop a model that predicts the impact of the M-pleat pattern on filter efficiency from first principles, we used a semiempirical approach. Flow resistance associated with the pleat pattern is inferred from clean-water scaling data and then applied to a model that predicts filtration efficiency as a function of membrane plugging.

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Clik here to view.The flux through a filter can be described using Darcy’s law (Equation 1), in which J is the flux, ΔP is the pressure differential across a filter, and R_t is the total resistance to flow. R_t includes flow resistance from the membrane (R_m), upstream and downstream supports (R_s), and filter housing (R_h) (Equation 2). Substituting Equation 2 into Equation 1 gives Equation 3.

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Figure 2: Components of total filtration resistance as a function of membrane plugging

For a plugging stream, R_m will increase with throughput while R_s will be unchanged (assuming that upstream and downstream supports do not plug). R_h will change with flow rate (predictably).

As a membrane plugs, resistance from the membrane becomes an increasingly larger fraction of the total resistance (Figure 2). R_s and R_h decrease relative to R_m as membrane plugging increases and the scaling factor increases with increasing filtrate volume (and degree of membrane plugging).

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Table 1: Ten-inch cartridges evaluated in this study

Materials and Methods
Membranes and Devices: Four high-area cartridges were evaluated in this study (Table 1). They included 10-inch high-area sterile high-capacity (SHC) (referred to as SHC-HA) and SHC standard-area (SHC-SA) cartridge filters (from MilliporeSigma). Those products contain the same types of membranes: one layer each of 0.5-µm and 0.2-µm polyethersulfone (PES) asymmetric membrane. Standard-area versions of those devices contain about 0.5 m² of effective filtration area. Two additional 10-inch cartridge filters from different manufacturers also were evaluated. For small-scale tests, 25-mm membrane discs were installed into OptiScale 25 devices (from MilliporeSigma), which contain 3.5 cm² of effective filtration area.

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Table 2:  List of challenge streams for throughput tests (PBS = polybutylene succinate)

Challenge Streams: Three challenge streams were used in this study (Table 2) to represent small, medium, and large particle sizes and particle-size distributions (Figure 3). The challenge streams were concentrated to achieve a high degree of plugging (>90% flux decay at <1,000 L/m² of filtrate) within about 30 minutes at the process conditions.

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Figure 3: Particle-size distributions of the challenge streams used in this study; particle sizing data were gathered using a Malvern MasterSizer particle size analyzer.

Test Method: Both OptiScale 25 devices and 10-inch cartridges were tested for clean-water permeability at 10 psi (690 mbar) and 21–25 °C. Following the water permeability test, throughput tests using one of the challenge streams were conducted at 10 psi (690 mbar). That throughput testing ran until the membrane permeability was reduced by at least 95% compared with the clean water permeability.

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Table 3: Cartridge-to-disc scaling factors for water permeability (LMH/psi)

Results and Discussion
Water permeability data were collected for each membrane and device type (Table 3). For water, scalability factors for three of the four filters tested were about 0.5. The flow resistance from the supports was about the same as that of the membrane, resulting in about a 50% loss in productivity compared with that of the OptiScale 25 device. In the fourth device (SHRp-HA, containing the tighter 0.1-μm membrane) the membrane was responsible for a higher fraction of the total filter resistance, resulting in a higher scaling factor.

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Figure 4: Throughput curves for SHC-HA challenged with soy peptone

Throughput was evaluated using multiple feed streams representing a wide range of particle-size distributions. The soy peptone (papainic digest of soymeal from MilliporeSigma) stream had a relatively small median particle size (about 0.2 µm) and relatively narrow particle-size distribution. This stream would be expected to foul membranes through an internal pore-plugging mechanism. The study found that for this stream, the initial flux-scaling factor was similar to that of water. But as the membrane fouled, the flux-scaling factor converged toward one (Figure 4). That is a result of the membrane becoming an increasingly larger fraction of the total filter resistance as the membrane fouls.

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Figure 5: Comparing data and model predictions for filtration of soy peptone with SHC-HA; (A) scaling factor over time; (B) 10-inch cartridge throughput over time.

We plotted scaling factor as a function of filtration time for the SHC-HA cartridge and the model prediction (Figure 5). Figure 6 shows throughput scaling factors for the cartridges tested in this study at 30 minutes filtration time (about 95–99% flux decay). For that stream (using 2 g/L soy peptone), scaling factors are all within about 15% of one (within the approximate measurement uncertainty).

The Hy-Soy T (Kerry) stream had a large particle size and wide distribution. Figure 7 shows throughput and flux decay curves for this stream, and Figure 8 shows scaling factors at 30 minutes. This stream showed initial flux scaling factors as expected (similar to water). But as the filters plugged, the scaling factors diverged away from one rather than converging. That result is explained by the larger particle size in the Hy-Soy T stream. These particles cannot enter the membrane pore structure and therefore accumulate at the membrane surface, forming a cake.