Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1
Sumith Ranil Wickramasinghe
 -- 2172 2022-04-24 06:15:41 |
2 format correct
Conner Chen
Meta information modification 2172 2022-04-25 10:50:22 |

Video Upload Options

Do you have a full video?
Send video materials Upload full video

Confirm

Are you sure to Delete?
Yes No
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Wickramasinghe, S. Virus Filter Foulants. Encyclopedia. Available online: https://encyclopedia.pub/entry/22203 (accessed on 14 May 2024).
Wickramasinghe S. Virus Filter Foulants. Encyclopedia. Available at: https://encyclopedia.pub/entry/22203. Accessed May 14, 2024.
Wickramasinghe, Sumith. "Virus Filter Foulants" Encyclopedia, https://encyclopedia.pub/entry/22203 (accessed May 14, 2024).
Wickramasinghe, S. (2022, April 24). Virus Filter Foulants. In Encyclopedia. https://encyclopedia.pub/entry/22203
Wickramasinghe, Sumith. "Virus Filter Foulants." Encyclopedia. Web. 24 April, 2022.
Virus Filter Foulants
Edit

The major classes of foulants in virus filtration. This includes irreversible and reversible product aggregates and minor product variants that differ in their charge or hydrophobicity. Product variants arise because mammalian cell-derived biotherapeutics are heterogeneous. The product is defined based on the production process and not on a single molecular species. Product variants with different post-translational modifications can have different hydrophobicity, charge, and conformations. If present, HCP, proteases, and nucleic acids can also foul the virus filter.

virus filtration

1. Monoclonal Antibody Aggregates

Aggregation is a typical occurrence with mAbs and other therapeutic proteins. Several pathways have been proposed to describe the aggregation of proteins. They include agglomeration of monomers in their native states, aggregation of conformationally altered or chemically modified monomers, nucleation, and surface-induced aggregation [1][2][3]. Significant attention has been placed on non-native monomer aggregation, since exposed hydrophobic moieties tend to self-associate [1]. Some surfactants, osmolytes, and chaotropes induce aggregation because they denature the monomeric product, exposing more of the hydrophobic core and distorting the surface charge distribution [4]. Physical and biochemical events can also induce product degradation through enzymatic and non-enzymatic processes such as shock, light, and oxidation [5].
Physical or chemical perturbations that put a strain on the native conformation of biotherapeutic proteins such as mAbs can result in clipping or aggregation [6]. Such conditions include the presence of chaotropic chemical species, pH swings [7], shock, mechanical stress, increased concentration, and large temperature fluctuation [1][8][9]. The size, charge, and hydrophobicity of a mAb aggregate will differ from that of the native structure.
Interfacial damage can also affect the stability of a product monomer, especially at the air–liquid interface, which induces nucleation and aggregation [10]. Surface tension and physical adsorption on solid surfaces also lead to conformational changes [1][11][12]. Freezing and thawing of a product induces more aggressive fouling of virus filters [13][14][15]. Freeze–thaw-induced aggregation is due to conformational changes at the ice–water interface and by freeze concentration [1][11][16].
Reversible aggregates are usually a precursor to nucleation [17], followed by irreversible aggregation as the aggregates increase in size [18][19]. As buffer ionic strength increases, electrostatic repulsion between the mAbs decreases, whereas hydrophobic attraction between the mAb increases, often leading to product aggregation [11]. Aggregation and precipitation occur most easily at the product’s isoelectric point (pI) due to reduced electrostatic repulsion between individual product molecules [20][21].

1.1. Reversible Aggregates

Aggregation can occur through different pathways, resulting in aggregates that are reversible or irreversible [1][4]. mAb oligomers such as dimers, trimers, and tetramers are typically reversible [1][2]. Reversible aggregates are known as soluble aggregates, and the associated product monomers are not significantly denatured. Soluble aggregates are caused by interactions between product molecules via hydrogen bonding, electrostatic, or van der Waals forces [4][22]. These soluble aggregates can foul virus filters if their size exceeds the 20 nm size cut-off of most parvovirus filters.
Rayfield et al. [23] investigated the impact of mAb properties on virus filter filterability and showed that aggregates bigger than 17 nm were correlated to the flux decline during virus filtration [13][23]. Monoclonal antibodies typically have a hydrodynamic diameter of 9–12 nm; thus, the small oligomers can be 20 nm to as much as 50 nm in diameter. Other studies have shown that freeze-thawing of mAbs may not cause aggregation in significant amounts detectable by size exclusion chromatography due to the relatively small diameters of potential aggregates formed [15][24][25].

1.2. Irreversible Aggregates

When product dimers and trimers undergo further aggregation, they attain a critical mass where the aggregate can no longer remain soluble. These large aggregates then precipitate out of the solution [1]. The precipitates become visible and show increased turbidity and cloudiness. These large aggregates are known as irreversible aggregates. Large, insoluble aggregates have an increased propensity to foul the separation-active layer during virus filtration. Barnard et al. investigated the principal foulant of freeze–thawed mAb solutions and found that the freeze–thaw process could induce the formation of large aggregates (>1 μm) [15]. The use of 0.1 or 0.22 µm pore size prefilters can mitigate virus filter fouling to some extent by removing these large aggregates.
Irreversible aggregation is prevalent with denatured product monomers [2][4]. Chemical degradation, such as oxidation and deamidation, alters the surface charge of product monomers and affects colloidal stability [2]. The irreversible aggregation of a product results in product loss, although even very low levels of aggregation (<1%) can cause a significant increase in filter fouling. Hawe et al. studied mAb aggregates formed during freeze–thaw- and heat-induced thermal stress [12]. Other studies show that heat denatures mAbs and leads to irreversible mAb aggregation [2][4].

2. Host Cell Proteins (HCP), Proteases, and Nucleic Acids

HCPs feature significantly in the downstream processing of protein-based therapeutics [26]. HCPs include proteins, enzymes, and co-enzymes which emanate from the host cell used for product expression [27]. It is essential to remove HCPs from therapeutic proteins because they can elicit an immune response. There are regulatory requirements for robust HCP removal before clinical trials of drug candidates to prevent the development of anti-CHO antibodies by volunteers [28][29]. Some HCPs can co-elute with the mAbs through polishing and purification steps, either due to binding to the resin or to association with the mAb product [27][30][31][32]. Zhang et al. identified over 500 HCPs in a cell culture sample and tracked their fate through downstream processing unit operations [33]. After studying nine different mAbs, they determined that actin and clusterin were most abundant in protein A eluates [33].
Enzymatic HCPs (host cell proteases) can clip or denature product monomers, expose hydrophobic residues and charged moieties, and alter the product’s biophysical properties. Denatured product monomers with exposed residues induce virus filter fouling by adsorptive processes in addition to mAb–mAb and mAb–HCP association. Host cell proteases have been reported to result in the fragmentation of mAb products, with increased susceptibility to nucleation and aggregation [34]. However, proteases themselves are probably not principal foulants of virus filters, since virus filtration occurs towards the end of downstream processing, where only trace amounts of non-mAb impurities may be detected [35][36].
HCP diminishes the biotherapeutic quality of biotherapeutic products and increases downstream processing costs. If HCPs are not sufficiently removed, they could potentially induce flux decay during virus filtration. HCPs have a range of biophysical properties, such as pI (2–11) and mass (10–200 kDa), which can be used to separate the HCP from the biotherapeutic [37][38]. Protein A chromatography significantly reduces HCPs in the mAb product due to high selectivity for the Fc region of mAbs [30]. Several studies reported that the propensity of different HCPs to bind and co-elute with mAbs from protein A columns vary from mAb to mAb [31][39]. Problematic HCPs are many and include lipoprotein lipase, nidogen-1, clusterin, histones, keratins, phospholipases, ribosomal proteins, and serine proteases [37].

3. Endotoxins

Endotoxins or lipopolysaccharides (LPS) are contaminants that can enter the process through growth media or other cell culture additives used in mammalian cell cultures. LPS are produced by Gram-negative bacteria, commonly used in recombinant DNA production [40][41][42]. Endotoxins are commonly found contaminants in mammalian cell-derived therapeutics [43]. LPS are complex molecular conjugates of an amphiphilic component (lipid A) and a polar polysaccharide component [41][42]. The isoelectric point of LPS ranges from 1 to 4 [38]. LPS removal techniques that have been reported include two-phase extraction, affinity chromatography, and ion exchange chromatography [42].
LPS have been reported to have a high affinity for some biotherapeutic proteins [43]. LPS and therapeutic proteins can form micellar aggregates, complicating the removal process and potentially carrying over into the virus filtration step [42][44]. Phosphorylated moieties of LPS electrostatically bind with the carboxyl moieties of amino acids in the biologic of interest [42][44]. Solutions of 0.5 M arginine have been shown to promote LPS clearance during polishing steps [44].
LPS have molecular masses ranging from 3–40 kDa, which vary due to their polysaccharide chain lengths [38][42]. Endotoxins can co-elute with mAbs and Fc-fusion proteins from protein A resins by molecular conjugation through hydrophobic and electrostatic interactions, ultimately causing problems during virus filtration. Endotoxin-contaminated mAb streams have an increased propensity to cause virus filter fouling. Removal of endotoxins through ion exchange polishing steps increases the virus filtration capacity of virus filters.

4. Product-Mediated Foulants

4.1. Charge Variants

The charge variant profile is a critical quality attribute of mAbs [45] and Fc-fusion proteins. Charge variants in mAbs can result from post-translational modifications (PTMs), such as deamidation of asparagine, C-terminal lysine variants, and glycosylation [46][47][48][49]. Glycans are mostly polar, hydrophilic oligosaccharides that can induce micro-differences in the surface charge of a glycoprotein. Negatively-charged glycans incorporating phosphorylated mannose and sialic acid can introduce micro-heterogeneities. Charge heterogeneity is observed in isoelectric-focusing electropherograms of most glycoproteins [46]. Acidic and basic variants of glycoproteins such as mAbs and Fc-fusion proteins can have different glycan profiles [50][51]. Meyer et al. [44] reported that specific charge variants of a mAb candidate were aggregation prone. Acidic variants of this mAb showed more pronounced hydrophobicity [52].
The net charge and surface charge distribution of glycoproteins change with buffer pH [48][53][54]. The pI of a protein is the pH value at which the net charge is zero [20]. For most mAbs, the pI ranges from 6.5–9.5 [20]. There is more biochemical variability with Fc-fusion proteins. A protein will be net negatively charged when the buffer pH is above the pI and positively charged when the buffer pH is below the pI [55]. Exposed surface residues on a glycoprotein can become protonated or deprotonated depending on the buffer pH, thereby inducing localized charged groups [56]. Charged moieties due to glycosylation, phosphorylation, and other PTMs affect the net charge of glycoproteins and their interactions with other product monomers and virus filtration membranes [55].

4.2. Denatured Variants

Hydrophobic interaction is the preferential association of non-polar residues in aqueous media [57]. Amino acids with non-polar side chains are typically hydrophobic, e.g., valine, leucine, proline, and tryptophan. Polar amino acids such as arginine impart hydrophilic attributes to glycoproteins [58]. When hydrophobic amino acids are surface-exposed on a glycoprotein, hydrophobicity increases. Hydrophobic amino acids tend to be buried in the globular core of most glycoproteins. The hydrophobicity of a protein is also affected by the buffer pH and the protein’s charge state [59]. When the buffer pH is close to the pI of the protein, the protein is the most hydrophobic [57]. Denaturation and unfolding of glycoproteins can lead to variants with a higher fouling propensity on virus filters.
The glycan appendages of glycoproteins also contribute to the final stable conformation, and glycan variation can introduce minor hydrophobicity variations. Careful handling and mild changes in formulation conditions will reduce the formation of conformational variants, which could foul virus filters or induce product aggregation.

4.3. Sequence Variants

Monoclonal antibodies and Fc-fusion proteins consist of amino acids in specific sequences that form secondary, tertiary, and quaternary structures. Sequence variants arise due to genetically unprogrammed amino acid substitutions, omissions, or insertions during biosynthesis [60]. Sequence variants result in macro-heterogeneities with biomolecular differences from the desired product [60]. Sequence variants possess different affinities to substrates [61] due to surface charge and hydrophobicity dissimilarities. The amino acid sequence of a glycoprotein determines its hydrophobicity, conformation, and charge, amongst other properties [60].
The primary structure (amino acid sequence) of a glycoprotein can determine intermolecular, monomeric association, and aggregation propensity [18]. Even minor sequence differences can cause conformational differences leading to product variants with different biophysical attributes and virus filter fouling propensity. Inadvertent substitution of hydrophilic amino acids with hydrophobic amino acids or vice versa in the polypeptide sequence amplifies sequence variants.

4.4. Micro-Heterogeneity-Induced Product Variants

mAbs, antibody fragments, bispecific antibodies, and Fc-fusion proteins are expressed in mammalian cells such as Chinese Hamster Ovary (CHO) cells for pharmaceutically relevant glycosylation profiles [62]. Flynn et al. reported that a typical CHO cell culture batch of mAbs has three major glycan species present, and they are G0F, G1F, and G2F [63]. These three dominant glycan structures are dependent on cell lineage and culture parameters [64]. E. coli expresses mostly insoluble, non-glycosylated variants [47]. Hybridomas offer a rapid expression template for initial product manufacture [62][65][66].
During cell culture and harvesting operations, expressed glycoproteins are usually not uniformly glycosylated [5][67][68]. Glycoproteins are expressed with a range of glycosylation profiles depending on cell culture conditions [69][70][71][72][73][74]. Micro-heterogeneity of glycoproteins can occur as a result of differences in glycosylation and other post-translational modifications. Variations in appended glycans introduce charge heterogeneity to the product monomer and determine the glycoprotein’s native fold state, aggregate susceptibility, and stability [75][76][77][78]. These product variants can affect the performance of virus filters.
Glycans are hydrophilic oligosaccharide moieties typically appended to glycoproteins in the cell during glycoprotein synthesis [79]. Glycans assist proper folding of the polypeptide chain before product secretion [46][67][80]. Most therapeutic proteins are glycoproteins. Glycoforms of protein products introduce structural heterogeneity, which affects their affinity to substrates, their stability, and other physicochemical characteristics of these therapeutic proteins [79][81][82]. Even in the same cell culture batch, a range of glycoforms occur [79][83][84]. Glycoforms occur due to skipped glycosylation sites on the glycoprotein or differences in the structure of appended glycans [67].
Glycan type and abundance can alter the product’s biophysical properties. Several studies have looked at the stability of different mAb glycoforms. These results show that aggregation is more prevalent in unglycosylated mAbs since glycans modulate aggregation [79][85]. Furthermore, a study showed that in terms of physical stability between pH 4–6, di-glycosylated IgG1-type mAbs were the most stable, and mono-glycosylated IgG1 was the least stable [86]. Post-translational modification can strongly affect the pI of a glycoprotein [46][79]. Variations in the pIs of product variants can affect hydrophobic and electrostatic interactions.
This entry is adapted from 10.3390/bioengineering9040155

References

  1. Manning, M.C.; Chou, D.K.; Murphy, B.M.; Payne, R.W.; Katayama, D.S. Stability of protein pharmaceuticals: An update. Pharm. Res. 2010, 27, 544–575.
  2. Philo, J.S.; Arakawa, T. Mechanisms of protein aggregation. Curr. Pharm. Biotechnol. 2009, 10, 348–351.
  3. Chakroun, N.; Hilton, D.; Ahmad, S.S.; Platt, G.W.; Dalby, P.A. Mapping the Aggregation Kinetics of a Therapeutic Antibody Fragment. Mol. Pharm. 2016, 13, 307–319.
  4. Woll, A.K.; Hubbuch, J. Investigation of the reversibility of freeze/thaw stress-induced protein instability using heat cycling as a function of different cryoprotectants. Bioprocess Biosyst. Eng. 2020, 43, 1309–1327.
  5. Dorai, H.; Ganguly, S. Mammalian cell-produced therapeutic proteins: Heterogeneity derived from protein degradation. Curr. Opin. Biotechnol. 2014, 30, 198–204.
  6. Diaz-Villanueva, J.F.; Diaz-Molina, R.; Garcia-Gonzalez, V. Protein Folding and Mechanisms of Proteostasis. Int. J. Mol. Sci. 2015, 16, 17193–17230.
  7. Sule, S.V.; Cheung, J.K.; Antochshuk, V.; Bhalla, A.S.; Narasimhan, C.; Blaisdell, S.; Shameem, M.; Tessier, P.M. Solution pH that minimizes self-association of three monoclonal antibodies is strongly dependent on ionic strength. Mol. Pharm. 2012, 9, 744–751.
  8. Zheng, J.Y.; Janis, L.J. Influence of pH, buffer species, and storage temperature on physicochemical stability of a humanized monoclonal antibody LA298. Int. J. Pharm. 2006, 308, 46–51.
  9. Salinas, B.A.; Sathish, H.A.; Shah, A.U.; Carpenter, J.F.; Randolph, T.W. Buffer-Dependent Fragmentation of a Humanized Full-Length Monoclonal Antibody. J. Pharm. Sci. 2010, 99, 2962–2974.
  10. Shieh, I.C.; Patel, A.R. Predicting the Agitation-Induced Aggregation of Monoclonal Antibodies Using Surface Tensiometry. Mol. Pharm. 2015, 12, 3184–3193.
  11. Kueltzo, L.A.; Wang, W.; Randolph, T.W.; Carpenter, J.F. Effects of solution conditions, processing parameters, and container materials on aggregation of a monoclonal antibody during freeze-thawing. J. Pharm. Sci. 2008, 97, 1801–1812.
  12. Hawe, A.; Kasper, J.C.; Friess, W.; Jiskoot, W. Structural properties of monoclonal antibody aggregates induced by freeze–thawing and thermal stress. Eur. J. Pharm. Sci. 2009, 38, 79–87.
  13. Wickramasinghe, S.R.; Namila; Fan, R.; Qian, X. Virus Removal and Virus Purification. In Current Trends and Future Developments on (Bio-) Membranes; Elsevier: Amsterdam, The Netherlands, 2019; pp. 69–96.
  14. Kern, G.; Krishnan, M. Virus Removal by Filtration: Points to Consider. BioPharm Int. 2006, 19, 32–41.
  15. Barnard, J.G.; Kahn, D.; Cetlin, D.; Randolph, T.W.; Carpenter, J.F. Investigations into the fouling mechanism of parvovirus filters during filtration of freeze-thawed mAb drug substance solutions. J. Pharm. Sci. 2014, 103, 890–899.
  16. Barnett, G.V.; Qi, W.; Amin, S.; Lewis, E.N.; Razinkov, V.I.; Kerwin, B.A.; Liu, Y.; Roberts, C.J. Structural Changes and Aggregation Mechanisms for Anti-Streptavidin IgG1 at Elevated Concentration. J. Phys. Chem. B 2015, 119, 15150–15163.
  17. Li, W.; Prabakaran, P.; Chen, W.; Zhu, Z.; Feng, Y.; Dimitrov, D.S. Antibody Aggregation: Insights from Sequence and Structure. Antibodies 2016, 5, 19.
  18. Wang, W. Protein aggregation and its inhibition in biopharmaceutics. Int. J. Pharm. 2005, 289, 1–30.
  19. Vazquez-Rey, M.; Lang, D.A. Aggregates in monoclonal antibody manufacturing processes. Biotechnol. Bioeng. 2011, 108, 1494–1508.
  20. Novák, P.; Havlíček, V. Protein Extraction and Precipitation. In Proteomic Profiling and Analytical Chemistry; Elsevier: Amsterdam, The Netherlands, 2016; pp. 51–62.
  21. Franco, R.; Daniela, G.; Fabrizio, M.; Ilaria, G.; Detlev, H. Influence of osmolarity and pH increase to achieve a reduction of monoclonal antibodies aggregates in a production process. Cytotechnology 1999, 29, 11–25.
  22. Leckband, D.; Israelachvili, J. Intermolecular forces in biology. Q. Rev. Biophys. 2001, 34, 105–267.
  23. Rayfield, W.J.; Roush, D.J.; Chmielowski, R.A.; Tugcu, N.; Barakat, S.; Cheung, J.K. Prediction of viral filtration performance of monoclonal antibodies based on biophysical properties of feed. Biotechnol. Prog. 2015, 31, 765–774.
  24. Barnard, J.G.; Singh, S.; Randolph, T.W.; Carpenter, J.F. Subvisible particle counting provides a sensitive method of detecting and quantifying aggregation of monoclonal antibody caused by freeze-thawing: Insights into the roles of particles in the protein aggregation pathway. J. Pharm. Sci. 2011, 100, 492–503.
  25. Bria, C.R.; Jones, J.; Charlesworth, A.; Ratanathanawongs Williams, S.K. Probing Submicron Aggregation Kinetics of an IgG Protein by Asymmetrical Flow Field-Flow Fractionation. J. Pharm. Sci. 2016, 105, 31–39.
  26. Namila, F.N.U.; Zhang, D.; Traylor, S.; Nguyen, T.; Singh, N.; Wickramasinghe, R.; Qian, X. The effects of buffer condition on the fouling behavior of MVM virus filtration of an Fc-fusion protein. Biotechnol. Bioeng. 2019, 116, 2621–2631.
  27. Aboulaich, N.; Chung, W.K.; Thompson, J.H.; Larkin, C.; Robbins, D.; Zhu, M. A novel approach to monitor clearance of host cell proteins associated with monoclonal antibodies. Biotechnol. Prog. 2014, 30, 1114–1124.
  28. Gutierrez, A.H.; Moise, L.; De Groot, A.S. Of and men: A new perspective on host cell proteins. Hum. Vaccin. Immunother. 2012, 8, 1172–1174.
  29. Doneanu, C.E.; Xenopoulos, A.; Fadgen, K.; Murphy, J.; Skilton, S.J.; Prentice, H.; Stapels, M.; Chen, W. Analysis of host-cell proteins in biotherapeutic proteins by comprehensive online two-dimensional liquid chromatography/mass spectrometry. MAbs 2012, 4, 24–44.
  30. Shukla, A.A.; Hinckley, P. Host cell protein clearance during protein a chromatography: Development of an improved column wash step. Biotechnol. Prog. 2008, 24, 1115–1121.
  31. Nogal, B.; Chhiba, K.; Emery, J.C. Select host cell proteins coelute with monoclonal antibodies in protein A chromatography. Biotechnol. Prog. 2012, 28, 454–458.
  32. Ahluwalia, D.; Dhillon, H.; Slaney, T.; Song, H.; Boux, H.; Mehta, S.; Zhang, L.; Valdez, A.; Krishnamurthy, G. Identification of a host cell protein impurity in therapeutic protein, P1. J. Pharm. Biomed. Anal. 2017, 141, 32–38.
  33. Zhang, Q.; Goetze, A.M.; Cui, H.; Wylie, J.; Trimble, S.; Hewig, A.; Flynn, G.C. Comprehensive tracking of host cell proteins during monoclonal antibody purifications using mass spectrometry. MAbs 2014, 6, 659–670.
  34. Gao, S.X.; Zhang, Y.; Stansberry-Perkins, K.; Buko, A.; Bai, S.; Nguyen, V.; Brader, M.L. Fragmentation of a highly purified monoclonal antibody attributed to residual CHO cell protease activity. Biotechnol. Bioeng. 2011, 108, 977–982.
  35. Bolton, G.R.; Spector, S.; LaCasse, D. Increasing the capacity of parvovirus-retentive membranes: Performance of the Viresolve™ Prefilter. Biotechnol. Appl. Biochem. 2006, 43, 55–63.
  36. Charcosset, C. 4—Virus filtration. In Membrane Processes in Biotechnology and Pharmaceutics; Charcosset, C., Ed.; Elsevier: Amsterdam, The Netherlands, 2012; pp. 143–167.
  37. Gilgunn, S.; El-Sabbahy, H.; Albrecht, S.; Gaikwad, M.; Corrigan, K.; Deakin, L.; Jellum, G.; Bones, J. Identification and tracking of problematic host cell proteins removed by a synthetic, highly functionalized nonwoven media in downstream bioprocessing of monoclonal antibodies. J. Chromatogr. A 2019, 1595, 28–38.
  38. Kornecki, M.; Mestmacker, F.; Zobel-Roos, S.; Heikaus de Figueiredo, L.; Schluter, H.; Strube, J. Host Cell Proteins in Biologics Manufacturing: The Good, the Bad, and the Ugly. Antibodies 2017, 6, 13.
  39. Luhrs, K.A.; Harris, D.A.; Summers, S.; Parseghian, M.H. Evicting hitchhiker antigens from purified antibodies. J. Chromatogr. B Anal. Technol. Biomed. Life Sci. 2009, 877, 1543–1552.
  40. Chen, R.H.; Huang, C.J.; Newton, B.S.; Ritter, G.; Old, L.J.; Batt, C.A. Factors affecting endotoxin removal from recombinant therapeutic proteins by anion exchange chromatography. Protein Expr. Purif. 2009, 64, 76–81.
  41. Zhang, M.; Zhang, L.; Cheng, L.-H.; Xu, K.; Xu, Q.-P.; Chen, H.-L.; Lai, J.-Y.; Tung, K.-L. Extracorporeal endotoxin removal by novel l-serine grafted PVDF membrane modules. J. Membr. Sci. 2012, 405–406, 104–112.
  42. Ongkudon, C.M.; Chew, J.H.; Liu, B.; Danquah, M.K. Chromatographic Removal of Endotoxins: A Bioprocess Engineer’s Perspective. ISRN Chromatogr. 2012, 2012, 649746.
  43. Petsch, D.; Anspach, F.B. Endotoxin removal from protein solutions. J. Biotechnol. 2000, 76, 97–119.
  44. Ritzén, U.; Rotticci-Mulder, J.; Strömberg, P.; Schmidt, S.R. Endotoxin reduction in monoclonal antibody preparations using arginine. J. Chromatogr. B 2007, 856, 343–347.
  45. Kahle, J.; Watzig, H. Determination of protein charge variants with (imaged) capillary isoelectric focusing and capillary zone electrophoresis. Electrophoresis 2018, 39, 2492–2511.
  46. Zhou, Q.; Park, S.-H.; Boucher, S.; Higgins, E.; Lee, K.; Edmunds, T. N-linked oligosaccharide analysis of glycoprotein bands from isoelectric focusing gels. Anal. Biochem. 2004, 335, 10–16.
  47. Wingfield, P.T. Overview of the purification of recombinant proteins. Curr. Protoc. Protein Sci. 2015, 80, 6.1.1–6.1.35.
  48. Vlasak, J.; Ionescu, R. Heterogeneity of monoclonal antibodies revealed by charge-sensitive methods. Curr. Pharm. Biotechnol. 2008, 9, 468–481.
  49. Kahle, J.; Zagst, H.; Wiesner, R.; Watzig, H. Comparative charge-based separation study with various capillary electrophoresis (CE) modes and cation exchange chromatography (CEX) for the analysis of monoclonal antibodies. J. Pharm. Biomed. Anal. 2019, 174, 460–470.
  50. Dai, J.; Lamp, J.; Xia, Q.; Zhang, Y. Capillary Isoelectric Focusing-Mass Spectrometry Method for the Separation and Online Characterization of Intact Monoclonal Antibody Charge Variants. Anal. Chem. 2018, 90, 2246–2254.
  51. Liu, H.; Ren, W.; Zong, L.; Zhang, J.; Wang, Y. Characterization of recombinant monoclonal antibody charge variants using WCX chromatography, icIEF and LC-MS/MS. Anal. Biochem. 2019, 564–565, 1–12.
  52. Meyer, R.M.; Berger, L.; Nerkamp, J.; Scheler, S.; Nehring, S.; Friess, W. Identification of Monoclonal Antibody Variants Involved in Aggregate Formation—Part 1: Charge Variants. Eur. J. Pharm. Biopharm. 2020, 158, 123–131.
  53. Rohani, M.M.; Zydney, A.L. Role of electrostatic interactions during protein ultrafiltration. Adv. Colloid. Interface Sci. 2010, 160, 40–48.
  54. Robinson, J.; Roush, D.; Cramer, S.M. The effect of pH on antibody retention in multimodal cation exchange chromatographic systems. J. Chromatogr. A 2020, 1617, 460838.
  55. Snyder, L.R.; Kirkland, J.J.; Dolan, J.W. Introduction to Modern Liquid Chromatography, 3rd ed.; Wiley-VCH: Hoboken, NJ, USA, 2010; pp. 584–618.
  56. Lehermayr, C.; Mahler, H.C.; Mader, K.; Fischer, S. Assessment of net charge and protein-protein interactions of different monoclonal antibodies. J. Pharm. Sci. 2011, 100, 2551–2562.
  57. Fekete, S.; Veuthey, J.L.; Beck, A.; Guillarme, D. Hydrophobic interaction chromatography for the characterization of monoclonal antibodies and related products. J. Pharm. Biomed. Anal. 2016, 130, 3–18.
  58. Burns, A.; Olszowy, P.; Ciborowski, P. Biomolecules. In Proteomic Profiling and Analytical Chemistry; Elsevier: Amsterdam, The Netherlands, 2016; pp. 7–24.
  59. Fekete, S.; Guillarme, D.; Sandra, P.; Sandra, K. Chromatographic, Electrophoretic, and Mass Spectrometric Methods for the Analytical Characterization of Protein Biopharmaceuticals. Anal. Chem. 2016, 88, 480–507.
  60. Borisov, O.V.; Alvarez, M.; Carroll, J.A.; Brown, P.W. Sequence Variants and Sequence Variant Analysis in Biotherapeutic Proteins. In State-of-the-Art and Emerging Technologies for Therapeutic Monoclonal Antibody Characterization Volume 2. Biopharmaceutical Characterization: The NISTmAb Case Study; American Chemical Society: Washington, DC, USA, 2015; Volume 1201, pp. 63–117.
  61. Hinkle, J.D.; D’Ippolito, R.A.; Panepinto, M.C.; Wang, W.H.; Bai, D.L.; Shabanowitz, J.; Hunt, D.F. Unambiguous Sequence Characterization of a Monoclonal Antibody in a Single Analysis Using a Nonspecific Immobilized Enzyme Reactor. Anal. Chem. 2019, 91, 13547–13554.
  62. Bramer, C.; Tunnermann, L.; Gonzalez Salcedo, A.; Reif, O.W.; Solle, D.; Scheper, T.; Beutel, S. Membrane Adsorber for the Fast Purification of a Monoclonal Antibody Using Protein A Chromatography. Membranes 2019, 9, 159.
  63. Flynn, G.C.; Chen, X.; Liu, Y.D.; Shah, B.; Zhang, Z. Naturally occurring glycan forms of human immunoglobulins G1 and G2. Mol. Immunol. 2010, 47, 2074–2082.
  64. Srebalus Barnes, C.A.; Lim, A. Applications of mass spectrometry for the structural characterization of recombinant protein pharmaceuticals. Mass Spectrom. Rev. 2007, 26, 370–388.
  65. Herschel, T.; El-Armouche, A.; Weber, S. Monoclonal antibodies, overview and outlook of a promising therapeutic option. Dtsch. Med. Wochenschr. 2016, 141, 1390–1394.
  66. Ribatti, D. Edelman’s view on the discovery of antibodies. Immunol. Lett. 2015, 164, 72–75.
  67. Dicker, M.; Strasser, R. Using glyco-engineering to produce therapeutic proteins. Expert Opin. Biol. 2015, 15, 1501–1516.
  68. Zydney, A.L. Perspectives on integrated continuous bioprocessing—opportunities and challenges. Curr. Opin. Chem. Eng. 2015, 10, 8–13.
  69. Rowe, L.; El Khoury, G.; Lowe, C.R. Affinity Chromatography: Historical and Prospective Overview. Biopharm. Prod. Technol. 2012, 1, 223–282.
  70. Wang, Y.; Li, X.; Liu, Y.H.; Richardson, D.; Li, H.; Shameem, M.; Yang, X. Simultaneous monitoring of oxidation, deamidation, isomerization, and glycosylation of monoclonal antibodies by liquid chromatography-mass spectrometry method with ultrafast tryptic digestion. MAbs 2016, 8, 1477–1486.
  71. Yang, X.; Kim, S.M.; Ruzanski, R.; Chen, Y.; Moses, S.; Ling, W.L.; Li, X.; Wang, S.C.; Li, H.; Ambrogelly, A.; et al. Ultrafast and high-throughput N-glycan analysis for monoclonal antibodies. MAbs 2016, 8, 706–717.
  72. Strohl, W.R.; Strohl, L.M. Therapeutic Antibody Engineering: Current and Future Advances Driving the Strongest Growth Area in the Pharmaceutical Industry; Elsevier Science & Technology: Cambridge, UK, 2012; Volume 11.
  73. Radhakrishnan, D.; Robinson, A.S.; Ogunnaike, B.A. Controlling the Glycosylation Profile in mAbs Using Time-Dependent Media Supplementation. Antibodies 2017, 7, 1.
  74. Ivarsson, M.; Villiger, T.K.; Morbidelli, M.; Soos, M. Evaluating the impact of cell culture process parameters on monoclonal antibody N-glycosylation. J. Biotechnol. 2014, 188, 88–96.
  75. Wu, W.; Song, H.; Slaney, T.; Ludwig, R.; Tao, L.; Das, T. Characterization of Protein Therapeutics by Mass Spectrometry. In Protein Analysis Using Mass Spectrometry: Accelerating Protein Biotherapeutics from Lab to Patient; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2017; pp. 221–249.
  76. Gagneux, P.; Varki, A. Evolutionary considerations in relating oligosaccharide diversity to biological function. Glycobiology 1999, 9, 747–755.
  77. Barinka, C.; Sacha, P.; Sklenar, J.; Man, P.; Bezouska, K.; Slusher, B.S.; Konvalinka, J. Identification of the N-glycosylation sites on glutamate carboxypeptidase II necessary for proteolytic activity. Protein Sci. 2004, 13, 1627–1635.
  78. Mrázek, H.; Weignerová, L.; Bojarová, P.; Novák, P.; Vaněk, O.; Bezouska, K. Carbohydrate synthesis and biosynthesis technologies for cracking of the glycan code: Recent advances. Biotechnol. Adv. 2012, 31, 17–37.
  79. Zhou, Q.; Qiu, H. The Mechanistic Impact of N-Glycosylation on Stability, Pharmacokinetics, and Immunogenicity of Therapeutic Proteins. J. Pharm. Sci. 2019, 108, 1366–1377.
  80. Wada, R.; Matsui, M.; Kawasaki, N. Influence of N-glycosylation on effector functions and thermal stability of glycoengineered IgG1 monoclonal antibody with homogeneous glycoforms. MAbs 2019, 11, 350–372.
  81. Planinc, A.; Bones, J.; Dejaegher, B.; Van Antwerpen, P.; Delporte, C. Glycan characterization of biopharmaceuticals: Updates and perspectives. Anal. Chim. Acta 2016, 921, 13–27.
  82. Sola, R.J.; Griebenow, K. Effects of glycosylation on the stability of protein pharmaceuticals. J. Pharm. Sci. 2009, 98, 1223–1245.
  83. Wang, Z.; Zhu, J.; Lu, H. Antibody glycosylation: Impact on antibody drug characteristics and quality control. Appl. Microbiol. Biotechnol. 2020, 104, 1905–1914.
  84. Liu, H.; Gaza-Bulseco, G.; Faldu, D.; Chumsae, C.; Sun, J. Heterogeneity of Monoclonal Antibodies. J. Pharm. Sci. 2008, 97, 2426–2447.
  85. Hari, S.B.; Lau, H.; Razinkov, V.I.; Chen, S.; Latypov, R.F. Acid-induced aggregation of human monoclonal IgG1 and IgG2: Molecular mechanism and the effect of solution composition. Biochemistry 2010, 49, 9328–9338.
  86. Alsenaidy, M.A.; Okbazghi, S.Z.; Kim, J.H.; Joshi, S.B.; Middaugh, C.R.; Tolbert, T.J.; Volkin, D.B. Physical stability comparisons of IgG1-Fc variants: Effects of N-glycosylation site occupancy and Asp/Gln residues at site Asn 297. J. Pharm. Sci. 2014, 103, 1613–1627.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
Upload a video for this entry
Information
Subjects: Engineering, Biomedical
Contributor MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
Sumith Wickramasinghe
View Times: 392
Entry Collection: Biopharmaceuticals Technology
Revisions: 2 times (View History)
Update Date: 25 Apr 2022
Table of Contents
    1000/1000
    Hot Most Recent
    Feedback