REVIEW
Aggregates in Monoclonal Antibody
Manufacturing Processes
Marı́a Vázquez-Rey,1 Dietmar A. Lang2
1
Manufacturing Science and Technology, Lonza Biologics Porriňo SL, A Relva s/n, 36410,
Porriño, Pontevedra, Spain
2
Technology Development, Lonza Biologics plc., 228 Bath Road, Slough, SL1 4DX, UK
Qiagen Manchester Ltd., Skelton House, Lloyd Street North, Manchester, M15 6SH, UK;
telephone: 0044-161-204-1161; fax: 0044-161-204-1200; e-mail: [email protected].
Received 5 December 2010; revision received 6 March 2011; accepted 30 March 2011
Published online 7 April 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/bit.23155
ABSTRACT: Monoclonal antibodies have proved to be a
highly successful class of therapeutic products. Large-scale
manufacturing of pharmaceutical antibodies is a complex
activity that requires considerable effort in both process and
analytical development. If a therapeutic protein cannot be
stabilized adequately, it will lose partially or totally its
therapeutic properties or even cause immunogenic reactions
thus potentially further endangering the patients’ health.
The phenomenon of protein aggregation is a common issue
that compromises the quality, safety, and efficacy of antibodies and can happen at different steps of the manufacturing process, including fermentation, purification, final
formulation, and storage. Aggregate levels in drug substance
and final drug product are a key factor when assessing
quality attributes of the molecule, since aggregation might
impact biological activity of the biopharmaceutical. In this
review it is analyzed how aggregates are formed during
monoclonal antibody industrial production, why they have
to be removed and the manufacturing process steps that are
designed to either minimize or remove aggregates in the final
product.
Biotechnol. Bioeng. 2011;108: 1494–1508.
ß 2011 Wiley Periodicals, Inc.
KEYWORDS: aggregates; monoclonal antibody; manufacturing processing
Introduction
Currently, therapy with monoclonal antibodies (mAb) is the
largest growth area in the pharmaceutical industry. The FDA
has approved 26 mAb for clinical use against cancer, Crohn’s
disease, rheumatoid arthritis, and antiviral prophylaxes
among other diseases (Birch and Racher, 2006; Scolnik,
Correspondence to: Dietmar A. Lang Qiagen Manchester Ltd., Skelton House, Lloyd
Street North, Manchester, M15 6SH, UK; telephone: 0044-161-204-1161; fax: 0044-161204-1200.
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2009; Wang et al., 2009) widely used in antitumor therapy
because of their specificity. mAbs are directed to their
tumoral targets (antigen) without affecting healthy tissues or
with minimal effects on them.
A standard mAb manufacturing process using mammalian cell culture is depicted in Figure 1. Cells from a cell bank
are thawed and cultured in small flasks. Over a period of
approximately 2 weeks, cells are grown in increasingly larger
volumes to provide a seed culture (inoculum) for the
fermentation tanks. Once cells are inoculated into the
production bioreactor (up to 20,000 L scale), they are grown
under controlled conditions to an optimal density for
maximum productivity. At the end of the production phase,
the harvest step separates the cells from the cell culture
supernatant to recover the product in the liquid phase. This
is typically done by either centrifugation and/or microfiltration (Sommerfeld and Strube, 2005).
The next step is in most cases a capture step using Protein
A chromatography and, since mAb elutes at low pH values, a
virus inactivation step follows at low pH. Additional process
steps include typically ion exchange chromatography
operations for polishing and/or impurities removal
(Faherner et al., 2001), in particular host cell proteins
(HCP), DNA, endotoxin, leached protein A, and aggregates.
In addition, membrane filtration plays an important role for
product concentration and buffer exchange. To guarantee
product safety, potential viral contaminants are removed by
filtration using nanofilters that has traditionally been
accepted as a robust method for virus clearance (Ray and
Tarrach, 2009). Once product impurities have been
removed and conditioning buffer has been added, product
is filled into bags, bottles, or stainless steel tanks for
subsequent storage until filled into vials.
Protein aggregation (oligomerization) behavior was first
studied in the 1960’s and was described by the simple
Lumry–Eyring model (Lumry and Eyring, 1954). Protein
aggregation is a common phenomenon during protein drug
ß 2011 Wiley Periodicals, Inc.
Figure 1. Outline of steps involved in a general mAb manufacturing process comprising (1) inoculum expansion, (2) product fermentation, (3) product primary recovery, (4)
product purification, (5) product formulation.
development but the mechanisms of aggregation are poorly
understood (Wang, 2005). Aggregates can initially exist as
small dimers or fragments and progress toward larger
structures, such as sub-visible or visible particles, if such a
transition becomes thermodynamically favorable (CórdobaRodrı́guez, 2008). Also, protein molecules may be unfolded
or partially unfolded, that is, the higher order structure of
the protein may be disrupted and, thereby, the protein’s
hydrophobic regions are exposed, promoting intermolecular interactions leading to aggregation or subsequently to
precipitation phenomena (Kiese et al., 2008).
Protein aggregates can be classified in several ways,
including
soluble/insoluble,
covalent/non-covalent,
reversible/non-reversible, and native/denatured according
to Cromwell et al. (2006). There is, however, no uniform
terminology for aggregate sizes and types so that Philo
(2006) introduced the classification: (1) rapidly reversible
non-covalent small oligomers (dimer, trimer, tetramer,
etc.); (2) irreversible non-covalent oligomers; (3) covalent
oligomers (e.g., disulfide-linked); (4) ‘‘large’’ aggregates
(>10 –mer), which could be reversible if non-covalent; (5)
‘‘very large’’ aggregates (diameter 50 nm to 3000 nm),
which could be reversible if non-covalent; and (6) visible
particulates (‘‘snow’’), which are probably irreversible.
Aggregation can arise from non-covalent interactions or
from covalently linked species (Manning et al., 2010).
Vázquez-Rey and Lang: Aggregates in mAb Manufacturing Processes
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Soluble aggregates have been defined as those that are not
visible as discrete particles and that may not be removed by a
0.22 mm filter (Cromwell et al., 2006). On the other hand,
insoluble aggregates are those that are often visible, and can
be removed either by 0.22 mm filtration or by centrifugation
at mild conditions (Knutson et al., 1979). Generally, they
can often be related to the precipitation of HCP impurities
rather than the product itself (Yigzaw et al., 2006).
Covalent aggregation is associated to chemical binding
between two or more monomers or with the chemical
linking of partially unfolded molecules with each other.
Disulfide bonds between previously unpaired free thiols are
a common mechanism for covalent aggregation (Andya
et al., 2003). Oxidation of tyrosine residues may also result
in covalent aggregation through the formation of bi-tyrosine
(Creed, 1984). For some proteins, a covalent interaction
between monomers is required to form a stable protein
structure.
A simplified model of non-covalent aggregation is given
in Mahler et al. (2005). Non-covalent aggregates are formed
when proteins associate and bind based on structural
regions of charge or polarity (Patel et al., 2011). Reversible
protein aggregation typically results from relatively weak
non-covalent protein interactions such as hydrophobic/
hydrophilic interactions with short distances between them.
The reversibility is sometimes indicative of the presence of
equilibrium between the monomer and higher molecular
weight protein variants. This equilibrium may shift as a
result of a change in solution conditions such as a decrease in
protein concentration or a change in pH. A weak, reversible
self-association of this type has been observed in a
monoclonal antibody to vascular endothelial growth factor
(VEGF; Moore et al., 1999). On some occasions, reversible
protein self-association produces an increase in the viscosity
of the protein solution (Liu et al., 2005).
The kinetics of protein aggregation has been extensively
investigated. Lumry and Eyring (1954) polymerization
model has been used as a starting point to study protein
aggregation. In the type of situation described by a Lumry–
Eyring model (Sánchez-Ruiz, 2010), the thermodynamically
stable protein with respect to unfolded and partially
unfolded states may undergo irreversible alteration processes that lead to some kind of a ‘‘final’’ state not being able
to fold back to the native one. One of the first models to
describe protein aggregation was developed based on the
understanding that changes in protein conformation can be
responsible for creating an altered state of the protein that is
then susceptible to aggregation (Lumry and Eyring, 1954).
Non-native aggregation can generally be considered as any
process which creates protein aggregates with secondary
structures at the monomer scale that are significantly
different from the dominant structures in the native state
(Roberts, 2007). The general model that explain the kinetics
of non-native aggregation considers that the protein (a
monomer fully folded) unfolds to form an unstable,
intermediate state that is in equilibrium with the native
protein structure. The unstable state serves as an
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intermediate for leading irreversibly to the aggregated state
(Andrews and Roberts, 2007; Powers and Powers, 2006).
This model has formed the basis of most subsequent
advances in understanding protein aggregation. Recently
Roberts (2007) has shown in more detail the general
understanding of aggregation as a multi-stage process and
how most available kinetic models of aggregation can be
grouped hierarchically in terms of which stage(s) they
include.
Recent reviews about aggregation kinetics (Morris et al.,
2009; Roberts, 2007; Sánchez-Ruiz, 2010) revealed that
protein aggregation is usually a higher order reaction due to
the fact that multiple conformational altered protein
molecules interact to create aggregated species. Despite this
general fact, situations in which the aggregation kinetics can
be of first order were also described (Kendrick et al.,
1998a,b) representing the case if the initial formation of
unfolding intermediates is rate limiting.
Protein heterogeneity also can be a contributing factor for
protein aggregation, as the probability of multiple protein
forms interacting with their environment is increased. In the
case of Epratuzumab, disulfide bond scrambling favored
covalent aggregate formation (Remmele et al., 2006).
Importance of Aggregate Removal
Differences in biological activity of the aggregates compared
to the activity of the monomeric protein can significantly
impair the potency of a protein-based drug. In such cases,
product efficacy may be compromised (Cromwell et al.,
2006). In general, a risk-based assessment of aggregates may
warrant specific studies which are helping to elucidate which
types of aggregates have a higher biological relevance. A
thorough analysis of the different conditions including
manufacture, storage, shipping, freeze and thaw cycles,
oxygen exposure, light, and physical stress to which the drug
is exposed from manufacturing until being administered,
can provide a rationale for special degradation studies such
as seeding or spiking of a specific aggregate or impurity into
a protein solution to observe the potential for further
aggregation (Córdoba-Rodrı́guez, 2008).
The extent to which aggregate levels impact the biological
activity of the protein is determined by multiple factors such
as molecular weight, structure, and solubility of aggregates
(Rosenberg, 2006). Individual susceptibility plays a key role,
whether determined by genetics or disease or a combination
of both (Singh, 2010). Protein aggregates pose risk in terms
of generation of immune responses to the therapeutic
protein product (Sharma, 2007). Of principal concern are
those immune responses associated with adverse clinical
effects such as neutralizing antibody that inhibits the efficacy
of the product (Franco et al., 1999) or, worse, crossreactivity neutralizes an endogenous protein counterpart,
causing severe hypersensitivity responses such as anaphylaxis (Rosenberg, 2006). Protein aggregates come in a variety
of forms such as fibrils (ordered aggregates), particulates
(irregular or spherical), skin, gels, or a combination of these.
The final form of aggregates seems to depend on the
aggregation pathway (Wang et al., 2010). The presence of
particulates in drug substances that are administered
intravenously has been shown to decrease microcirculation
as a consequence of the mechanical blockage of capillaries
(Lehr et al., 2002). Preclinical studies with properly designed
models may be useful to rank the factors as to their potential
for impacting immunogenicity (Singh, 2010). A first animal
model—transgenic mice—already showed that protein
aggregates are able to break the immune tolerance for a
protein—interferon beta. The potency of the aggregates to
break tolerance not only depends on aggregate percentage
but also largely on their physical properties such as degree of
denaturation, molecular orientation, and size (Van Beers
et al., 2010). Further investigations into what types of
aggregates are immunogenic, which protein types are most
prone to immune problems, what dosing is required and
how well do results from animal models predict effects in
humans are subjects for future research.
There is no consensus on the maximum allowable
aggregate levels in protein-based pharmaceutical products
because some proteins may be largely stable and safe despite
certain levels of aggregates, while for other proteins very
small changes in aggregate levels may significantly affect
protein stability, and even safety (Córdoba-Rodrı́guez, 2008;
Cromwell et al., 2006). The only group of aggregates that has
maximum allowable limits based on United States
Pharmacopeia,
USP
<788>
(also
European
Pharmacopoeia [Ph. Eur.] 2.9.19 and Ph. Eur. 2.9.20), is
the group of sub-visible particles. Solutions for injection
must be clear and practically free from particles. Limits for
soluble protein aggregates have to be set on a case-by-case
basis as there are no regulatory limits for aggregates in biotherapeutic preparations (Mahler et al., 2009). The
acceptance criteria for aggregates levels should reflect values
that maintain the safety and efficacy of the product (ICH.
Q6B Specifications: Test Procedures and Acceptance
Criteria for Biotechnological/Biological Products, 1999).
In the absence of information on clinical relevance and
process control, many specifications are instituted with
narrower than necessary acceptance ranges based on
manufacturing experience (Kozlowski and Swann, 2006).
The European Pharmacopoeia (2006a) specifies in the
monograph parenteral preparations, the requirements for
sterile solutions administered by injection like protein
solutions. Solutions for injection must be clear and
practically free from particles. They have to comply with
the test for sterility, the test for particulate contamination:
sub-visible particles, test for uniformity of content, and the
test for bacterial endotoxins or pyrogens. As protein
degradation often results in aggregation and precipitation,
particles in the solution present besides other points a
critical aspect.
The test for visible particles according to the European
Pharmacopoeia (2006b) describes a simple procedure for a
visual inspection of parenteral solutions. The aim is to assess
the quality of the solution in respect to particulate
contamination consisting of mobile undissolved particles
other than gas bubbles. The visual inspection is performed
with the help of a viewing station consisting of a matt black
and a non-glare white panel in vertical position next to each
other and a suitable white-light source. Non-labeled containers that are clean and dry on the outside are inspected for
particles by gently swirling and observing 5 s in front of the
white panel and in front of the black panel. As it is a nondestructive method a 100% control can be performed.
The test for sub-visible particles according to the European
Pharmacopoeia (2006c), which has to be performed for
solutions for injection, is conducted using the light
obscuration particle count test. The method allows a
determination of a size distribution using the principle of
light blockage. A suitable instrument calibrated with
spherical particles of known size between 10 and 25 mm is
used to examine a statistically relevant number of test
specimens after sample preparation. Solutions for injection
with a nominal volume of equal or less than 100 mL comply
with the test if the average number of particles in the tested
units does not exceed 6,000 per container equal to or grater
than 10 mm and 600 per container equal to or greater than
25 mm. Solutions with a nominal volume of more than 100 mL
comply with the test if the average number of particles in the
tested samples does not exceed 25/mL equal to or greater than
10 mm and 3/mL equal to or greater than 25 mm. The
microscopic particle count test is available as a second backup method in the European Pharmacopoeia (2006c).
Aggregates Formation During Monoclonal
Antibody Manufacturing Processes
There are many environmental factors that can lead to
aggregation (Remmele et al., 2006). Environmental conditions of production process such as temperature, protein
concentration, pH, oxygen, shear forces, and the ionic
strength may affect the amount of aggregate observed. The
presence of certain ligands, including specific ions, may
enhance aggregation. Stresses to the protein such as freezing,
exposure to air, or interactions with metal surfaces may
result in undesired post-translational molecule modification
even molecule unfolding, which then leads to the formation
of aggregates. Finally, mechanical stresses may cause protein
aggregation. Each of these environmental factors is typically
encountered during bioprocessing (Patel et al., 2011; Wang,
2005; Wang et al., 2010).
Aggregation During Cell Culture
Protein aggregation during cell culture is a widely reported
phenomenon and, in fact, aggregate levels up to 30% have
been reported for some mAbs in mammalian cell culture
(Kramarczyk et al., 2008). During cell culture, protein
aggregates might be formed (i) within the cell following
protein expression and (ii) once the protein is secreted into
Vázquez-Rey and Lang: Aggregates in mAb Manufacturing Processes
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1497
the cell culture medium (Cromwell et al., 2006). During
expression, accumulation of high amounts of protein may
lead to intracellular aggregation owing to either the interactions of unfolded protein molecules or to inefficient
recognition of the nascent peptide chain by molecular
chaperones responsible for proper folding (Zhang et al., 2004).
It is possible to influence the amount of aggregates produced during the cell culture process by carefully selecting the
optimal cell line and optimizing cell culture conditions such
as media components that will impact media osmolality and
conductivity, feed strategy, temperature, and pH (Gabrielson
et al., 2007). Mice cells cultured in a hollow fiber bioreactor,
exhibited a lower amount of aggregates in the protein secreted
when media pH and osmolarity were increased (Franco
et al., 1999). Cell culture temperature influence in aggregate
formation during cell culture was investigated by Cromwell
et al. (2006). They have reported that the greater the length of
time the protein was held in the cell culture medium at
elevated temperature, the greater the amount of aggregates
observed (Cromwell et al., 2006).
Secretion of the protein into the cell culture medium
exposes the protein to environmental stress such as
unfavorable pH conditions that may destabilize the protein.
Franco et al. (1999) studied a mAb with an isoelectric point
(pI) of pH 6.1–6.5 that was secreted into a cell culture
medium at pH 7.1–7.2. At the medium pH, the mAb
theoretically carried an overall negative charge, but there are
still some positive charges on the molecule. The electrostatic
forces between positive and negative charges on other mAb
molecules favor the formation of aggregates in solution. To
limit this phenomenon Franco et al. (1999) increased the pH
to such a degree that the net charge of the antibody should
be sufficient to effect repulsion between the molecules in
solution. Furthermore, to enhance the solubility, they
increased the ionic strength of the medium by adding a
solution of sodium chloride (98 g/L) to achieve an
osmolarity value of 350 mOsm/kg H2O (Franco et al., 1999).
Aggregation During Purification
When a therapeutic protein has been produced, it has to be
purified in order to reduce or eliminate any viruses,
aggregates, DNA (nucleotides in general), HCP, and other
process-related impurities. Protein purification processes
comprise multiple steps where different techniques are
employed, based on different separation principles such as
affinity, charge, size, hydrophobicity, or other properties of
the target protein compared to impurities (Sommerfeld and
Strube, 2005). As mentioned above, the result may be that
the protein experiences a wide range of pH, ionic strength,
protein concentrations, and contact materials during the
process. In addition to that, it has to be considered that at
large scale the protein is exposed to mechanical stresses such
as agitation in tanks and pumping (Thomas and Geer,
2011). Once the protein has been purified, it is typically filled
into bottles, bags, or cryo-vessels for further freezing and
storage. Each condition experienced by the protein may
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affect the degree of aggregation observed. In this section, the
effect of the above mentioned conditions on protein
aggregation will be discussed.
Exposure to Low pH Condition
Protein A chromatography is the capture step of choice for
purifying mAb after harvest (Faherner et al., 2001). It was
shown that a highly conserved histidyl residue in the center
of the Protein A binding site of IgGs faces a complementary
histidyl residue on Protein A (Chen, 1992). These residues
have a positive charge at low pH, thus repelling each other
and dissociating the Protein A–IgG hydrophilic interaction.
As a result, operating conditions for Protein A columns
typically require the use of low pH conditions (between pH 3
and 4). At this low pH proteins might undergo structural
changes that could contribute to product aggregation
(Chen, 1992; Krishnamurthy and Manning, 2002). In the
preparation of CamPath-1H1 (Alemtuzumab), which
elutes from the Protein A column with 0.1 M Sodium
Citrate, pH 3.2, the eluate contained approximately 25%
aggregated mAb (Phillips et al., 2001). Strategies to
minimize aggregate formation during Protein A chromatography have been considered and include the use of urea—a
chaotropic agent—at moderate concentrations (<2 M) as
an effective stabilizer or to carry out the separation under
low temperature conditions (Chen, 1992).
Low pH conditions are favorable purification conditions
for Protein A chromatography as well as for the viral
inactivation step. The viral inactivation step is intended to
modify the surface chemistry of viruses to render them
inactive or denatured. Low pH treatment has been shown to
successfully inactivate retroviruses for a variety of biotechnology products but also originates aggregates in the protein
pool due to the low pH exposure (Doms et al., 1985; Sofer,
2002). The following manufacturing step requires the
neutralization of the solution which supports stabilizing
the product. The use of strong bases (such as sodium
hydroxide) is avoided despite the advantage of low volume
addition due to the risk of product denaturation in the
localized region where the solution is added. Due to that the
use of higher concentrations of weaker bases (e.g., Tris base
solution) is preferred (Shukla et al., 2007). Some mAb
solutions might exhibit a turbid appearance following
neutralization. (Shukla et al., 2007).
Exposure to pH Conditions Near Protein Isoelectric Point
Proteins have ionizable groups such as carboxyl groups and
amino groups. Since the charge of these groups depends
on pH, a protein molecule depending on its pI may have
different charges at different pH values during manufacturing (Xia, 2007).
When a protein structure equally includes both positively
and negatively charged groups (i.e., at pH values close to the
pI), anisotropic charge distribution on the protein surface
can create dipoles. In such cases, protein–protein interactions are frequent, making assembly processes such as
aggregation energetically favorable (Striolo et al., 2002).
Agitation
Agitation is the mechanism used to achieve homogeneity of
protein pools during mAb purification manufacturing
processes. Mixing is especially critical in those purification
stages where buffers or solutions are added to product
solutions to adjust pH and/or conductivity for the next
separation steps. During agitation of protein solutions a gas/
liquid interface is created at which aggregation predominantly occurs. There have now been many studies
specifically investigating how proteins behave at gas–liquid
interfaces (Thomas and Geer, 2011).
Mahler et al. (2005) claimed that agitation in the presence
of a (hydrophobic) gas–liquid interface caused aggregation
of immunoglobulin-G1 (IgG1). Antibodies such as IgG1,
might also be susceptible to damage at air–liquid interfaces.
Harrison et al. (1998) showed that a recombinant scFV
antibody fragment suffered a first order loss of activity in a
partially filled, agitated vessel.
During agitation insoluble aggregates are released into
bulk solution (Carpenter et al., 1999; Fesinmeyer et al.,
2009). Fesinmeyer et al. (2009) reported that agitation of
70 mg/mL mAb solutions for a period of 65 h resulted in
increased solution turbidity. If stored statically, the solutions
became clear, with a layer of protein aggregate at the bottom,
consistent with the formation of insoluble aggregate
particles (Fesinmeyer et al., 2009).
Protein aggregation is commonly a second- or higherorder process. It is expected to increase with higher protein
concentration, however, Treuheit et al. (2002) has shown
that aggregation decreased with higher protein concentration if induced as a result of agitation (Treuheit et al., 2002).
This unexpected result may be explained by the rate-limiting
effect on aggregation of the air/water interface and the
critical nature of the air/water interface to protein ratio that
is greatest with decreased protein concentration (Treuheit
et al., 2002).
Apart from agitation, stirring and shaking are other
methods used to achieve homogeneity during mAb
production. These methods are used at large scale to
homogenize bottle contents, especially during final filling
operations. Mechanical stress associated to stirring and
shaking results in the formation of different species, sizes,
and amounts of non-covalent aggregates (Kiese et al., 2008).
Stirring has been reported to yield many insoluble, visible,
and sub-visible particles and high turbidity, whilst shaking is
reported to induce higher amounts of soluble aggregates
(Kiese et al., 2008).
Buffer Characteristics
During manufacturing, mAbs are suspended in various
buffer solutions. Salts and buffers have complex effects on
protein stability. Depending on the type and concentration
of the salt, the charged groups of the protein and the type of
ionic interactions between them, salts may have a stabilizing,
a destabilizing, or no effect (Kendrich et al., 2002). In this
section, the influence of buffers composition on aggregates
formation is explained.
Kameoka et al. (2007) have reported that the aggregation
effect of buffer species is associated to the specific molecular
interaction between buffer and IgG. It was indicated that the
specific interaction between buffer molecules (like citrate or
phosphate ions) and the Fc domain of IgG is the leading
mechanism in the aggregation of IgG (Kameoka et al., 2007).
Surfactants, such as polysorbates, are frequently added to
pharmaceutical protein solutions to mitigate the risk of
agitation-induced aggregate formation as they interfere with
hydrophobic interactions at the gas/liquid interface
(Fesinmeyer et al., 2009). Although surfactants are typically
used to minimize aggregate formation, for some proteins
like recombinant human growth hormone (rhGH), surfactants bind more strongly to the native state and they
decrease the free energy of denaturation which could
contribute to product degradation (Bam et al., 1998).
Antimicrobial preservatives, such as benzyl alcohol and
phenol, are often used in protein liquid formulations to
ensure sterility in multi-use products where by necessity the
container-closure must be breached multiple times in the
process of taking doses from the vial. Although the use of
this preservatives is not very extended because most of the
mAb are not multi-dose product, the preservatives (if
presents) often induce aggregation of proteins in aqueous
solution. For example, preservatives (e.g., phenol, m-cresol,
and benzyl alcohol) have been shown to induce aggregation
of granulocyte colony stimulating factor (Chi et al., 2003) or
recombinant interleukin-1 receptor (Remmele et al., 1998).
Since the contribution to solution ionic strength increases
with the valence of an ion, the use of polyvalent salts could
present significant challenges for the development of
aggregate-free protein therapeutic products (Fesinmeyer
et al., 2009). When the electrostatic coupling strength is
increased by increasing valence or lowering the dielectric
permits, this is accompanied by a strong accumulation of
counter ions close to the protein. Increasing salt concentration will induce protein aggregation and depending on the
ion valence less—trivalent—or far more—monovalent—
salt is required to suppress repulsion (Lund and Jonsson,
2003).
Ions resulting from salt dissolution can have a deleterious
effect on protein stability and promote aggregate formation.
A clear correlation between increased ionic strength and
increased aggregation was observed by Fesinmeyer et al.
(2009) and Gokarn et al. (2008), however, and as mentioned
at the beginning of this section, high concentrations of salt as
observed in other proteins can stabilize proteins through
the preferential exclusion mechanism. The effect correlates
with the Hofmeister series for anions: citrate3 /citrate2 >
PO34 HPO24 SO24 > OAc , F > Cl > Br > I >
ClO4 (Kendrick et al., 2002).
Vázquez-Rey and Lang: Aggregates in mAb Manufacturing Processes
Biotechnology and Bioengineering
1499
Equipment Contact Materials
A general review of the mechanisms of protein adsorption
on a solid surface is to be found in Nakanishi et al. (2001).
Stainless steel, a ubiquitous surface in bioprocessing, has
been shown to cause aggregation of mAbs. Two different
IgG4 mAbs were found to aggregate according to first-order
kinetics when exposed to stainless steel under high shear
conditions (Biddlecombe et al., 2007). In a different study,
exposure of an IgG to stainless steel particulates caused the
generation of much larger particles even when formulated
with a non-ionic surfactant (Tyagi et al., 2009). Bee et al.
(2009a) found that stainless steel induce aggregation of a
mAb with a second-order dependence on the steel surface
area and a zero-order dependence on the mAb concentration (Bee et al., 2009a). These studies demonstrate that
stainless steel microparticles can cause the aggregation of a
mAb. Also, the solution conditions are a critical parameter
for aggregates formation when mAbs are in contact with
stainless steel. Fe ions leached from steel have been reported
to cause oxidation of proteins resulting in aggregation (Lam
et al., 1997; Wang et al., 2007).
Stainless steel is a much extended construction material in
large-scale equipment used to manufacture mAbs, however,
along the processes other materials such as Teflon, glass, or
titanium can be found. These construction materials also
can lead to aggregates formation. A broad study of the
adsorption of 18 proteins on a titanium oxide can be found
in Imamura et al. (2008). Sluzky et al. (1992) studied the
aggregation of insulin in hydrophobic solid surfaces such as
Teflon. The monomer was denatured at the surface followed
by formation of microaggregates. Colombie et al. (2001)
suggested that polytetrafluoroethylene (PTFE) was much
more effective at causing aggregation of lysozyme than glass,
which confirms the view that hydrophobicity is the
governing factor in these phenomena.
Ultrafiltration
Ultrafiltration (UF) has been widely adopted as the method
of choice for protein formulation in the biotechnology
industry (Van Reis and Zydney, 2001). This purification step
is typically performed to exchange the buffer, reduce
conductivity, and to increase the protein concentration in
solution (Kiefhaber et al., 1991; Shire et al., 2004). However,
there are many challenges in applying UF to high
concentration protein solutions including product solubility, restrictive bulk and mass transport of protein solutions,
and product losses during recovery.
During UF, proteins are exposed to physical stress due to
pumping, with a typical process requiring at least 50 passes
through the pump (Harris et al., 2004). This stress may
result in an increase in protein aggregation (Van Reis et al.,
1997). Multiple passes through valves and the concomitant
microcavitation and air bubble entrainment, rather than
shear, have also been cited as a cause of aggregation during
filtration (Narendranathan and Dunnill, 1982).
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The extensive contact to membrane surfaces during UF is
another factor that contributes to aggregates formation. The
concentration of protein at the membrane surface may be
much higher than that of the bulk solution. This phenomenon
may lead to membrane fouling and to boundary layer polarization near the membrane which are considered to promote
aggregation (Bodalo et al., 2004; Kiefhaber et al., 1991).
Pumping
During downstream processing of mAb, the use of pumps is
quite extensive, especially in those steps that require flow
control. Typically lobular pumps are used to load product
onto chromatography units, during viral filtration steps, UF,
and during final filling (Gomme et al., 2006a; Thomas and
Geer, 2011). It is quite extended the knowledge that these
pumping processes expose mAb to mechanical shear forces
that might result into aggregates formation because most
damage to proteins during processing will increase with
higher fluid flow rates, for example, in passage through a
pump, however, it has not always been clear that any effects
are truly attributable to shear in the fluid mechanical sense,
which is caused by velocity gradients in moving liquids
(Thomas and Geer, 2011).
Cavitation is an additional pumping stress that aggressively creates and destroys microbubbles inside pumps and
valves (Bee et al., 2009b; Van Reis and Zydney, 2007).
During rotary lobe pumping of human albumin, a pump
with larger self-lubricated clearances caused more aggregation than a pump with smaller clearances (Gomme et al.,
2006b). This was attributed to ‘‘gap cavitation’’ occurring
and causing aggregation (Gomme et al., 2006b; Neumaier,
2000). Furthermore, there are studies that support that shear
stress is the factor that less influences aggregate formation
during pumping. Aggregation observed during the production of protein therapeutics is, therefore, not likely caused by
shear per se, but rather by the gas/liquid interface that is
created, exposure to solid surfaces, contamination by
particulates, or pump cavitation that is often associated
with shear (Bee et al., 2009b).
The type of pump to be used during manufacturing is
another factor influencing protein aggregation behavior.
Meireles et al. (1991) studied the effects of different pump
heads and observed an increase in turbidity of an albumin
preparation with pumping time at room temperature by
using a screw pump (Meireles et al., 1991). Likewise, it was
observed that the use of a peristaltic pump enhances
aggregate formation (Chandavarkar, 1990). In a recent
study, strong evidence was found of aggregation of viruslike
particles adsorbed to an adjuvant during recirculation
studies involving a peristaltic pump. In these studies,
increased back pressure in scaled down equipment demonstrated a proportional increase in aggregation, as measured
by light scattering. While this result was initially attributed
to pump ‘‘shear’’ increasing as a result of higher back
pressures, additional studies showed that if a different
tubing material was used, the levels of aggregation were
independent of back pressure. This result is interesting as
peristaltic pumps are frequently advertised as a mechanism
to minimize shear during processing. Thus, protein
aggregation via a peristaltic pump is unlikely related to
hydrodynamic shear mechanisms and requires further
investigation (Thomas and Geer, 2011).
Final Filling
Product filling is the final step in the manufacturing
process—no further purification after the fill. It is critical,
therefore, that the formation of aggregates during this step is
minimized if not prevented completely. For this reason it is
essential that compatibility of the protein formulation with
the filling equipment is assessed before production.
Fill-and-finish operations may use pumps that can
mechanically denature the protein because of shear stress
or introduce impurities that serve as nucleation sources of
protein aggregates (Tyagi et al., 2009). Some pistondisplacement pumps, for example, can interact with protein
drug product in a similar way that a car motor engine piston
interacts with lubricant oil. The intimate contact between
protein drug product and a piston rod can disrupt an
otherwise stable drug product. This was the case for an
antibody drug product that was found to have increased
levels of aggregate particles with more pump passes, as
determined by absorbance and light obscuration methods
(Cromwell et al., 2006).
Materials used during final filling operations also impact
aggregates formation. Glass from vials, rubber from
stoppers, silicone from stoppers, and syringes and tungsten
from syringes are some of the foreign particles which will
come into contact with drug product during final filling
operations (Carpenter et al., 1999). Many of these foreign
particles are electrostatically charged and, therefore, have the
potential to interact with proteins, protein aggregates, and
protein aggregate precursors to form heteronuclei. Such was
the case with prefilled syringes containing tungsten particles
shed during syringe barrel manufacturing, which served as
nuclei for aggregate formation (Eckhardt et al., 1991).
Syringes, stoppers, and other surfaces are treated with
silicone oil for lubrication or to inhibit protein binding but
silicone oil can actually induce protein aggregation (Jones
et al., 2005; Thirumangalathu et al., 2009). Substituting silicone
oil with other substances like Teflon1 has proven to cause
protein aggregation due to adsorption of protein molecules at
the solid/liquid interface as well (Sluzky et al., 1991).
Freeze, Thaw, and Storage
Most therapeutic proteins are delivered to patients in vials
that will be accidentally or deliberately shaken between
filling and delivery. Kiese et al. (2010) showed that shaking
vials cause IgG1 aggregation but that the insoluble
aggregates caused by the former could reversibly dissociate
into soluble aggregates. This aggregation was probably due
to the air–liquid interface in the vials.
Therapeutic proteins are usually frozen for long-term
storage of bulk drug substance prior to fill and finish
operations. Frozen storage of drug products is generally
preferred over liquid storage for several reasons including
increased product stability and shelf life, decreased
microbial growth, and elimination of foaming during
transport (Webb et al., 2002).
Freeze–thawing can be considered as a combination of
various stresses, like cold denaturation, by introducing ice–
liquid interfaces (Harding et al., 1999; Strambini and
Gabellieri, 1996), and by freeze-concentration (cryo-concentration) of solutes when the water crystallizes (Franks,
1982). This stress can also lead to pH shifts if the buffer salts
crystallize (Hawe et al., 2009; Van den Berg, 1996). Freezeconcentration can cause phase separation of excipients and
loss of native protein structure during subsequent drying
(Heller et al., 1997). All these stresses might cause protein
aggregation as reported for antibody freeze–thawing with
decreasing pH, which correlated well with Tm (protein
melting temperature) values (Kohle and Badkar, 2010;
Kueltzo et al., 2008; Manning et al., 2010). Partial unfolding
of proteins at the ice–freeze concentrate interface was found
to be one mechanism of aggregates formation during
freezing (Chang et al., 1996; Eckhardt et al., 1991; Strambini
and Gabellieri, 1996). Studies performed with human
growth hormone have also suggested that protein aggregation could be correlated with the area of the solid/liquid
interface (Eckhardt et al., 1991).
Low temperature can cause spontaneous unfolding of
proteins. Refolding and aggregation pathways compete for
unfolded protein molecules. Slow freezing of solutions can
lead to elevated concentrations of macromolecules interacting
over extended periods of time, increasing the likelihood that
an unfolded molecule will aggregate (Strambini and Gabellieri,
1996). On the contrary rapid freezing may lead to greater
concentration of solution polarization. Keeping in mind the
content and type of salt, and the damaging effect of the ice
surface area, an optimal freezing rate as function of protein,
and temperature would balance against the total exposure
time with ice surfaces present. Aggregation after freeze–
thawing was detected primarily by the formation of particles
in the size range from 1 to 25 mm, as well as aggregates of
about 0.5 mm (Kueltzo et al., 2008).
Heating and freeze–thawing of monoclonal IgG1 antibody formulations results in the formation of aggregates,
which differ considerably in size and structure (Bhatnagar
et al., 2007; Hawe et al., 2009; Vermeer and Norde, 2000).
Heating stress is mainly evidenced by the formation of small
soluble IgG aggregates in the size range up to 30 nm, which
are structurally changed. Freeze–thawing on the other hand
mainly leads to the formation of particles in the size range
from 1 to 25 mm and few aggregates in the size range of
500 nm (Hawe et al., 2009).
The containers commonly used for storage of bulk
solutions are polypropylene, polyethylene, glass vials,
Teflon1 and ethylene vinyl acetate (EVA) bioprocessing
bags. Kueltzo et al. (2008) demonstrated that less soluble
Vázquez-Rey and Lang: Aggregates in mAb Manufacturing Processes
Biotechnology and Bioengineering
1501
aggregate was detected at low pH (4) in the EVA bioprocessing bag and Teflon1 vials than in polypropylene tubes.
The preparation of freeze-dried mAb formulated without
excipients resulted in reversible solid-state protein structural
alteration. The mechanisms by which sugars/polyols can
improve the stability of a protein during drying and storage
are still incompletely understood. There are two main
hypotheses advanced to rationalize the role of solutes in
stabilizing proteins during drying and storage. One is the
‘‘glass dynamics hypothesis’’ and another is the ‘‘water
substitute hypothesis’’ (Chang et al., 2005). The ‘‘glass
dynamics hypothesis’’ states that a good stabilizer forms a
rigid, inert matrix into which the protein is molecularly
dispersed and can couple the motion of protein to the
motion of matrix. Dilution of proteins in the glass matrix
separates the protein molecules, and limited mobility in the
glass minimizes bimolecular interactions. Stabilization is via
a purely kinetic mechanism, and the stability of the protein
would be expected to correlate with the molecular mobility
in this rigid matrix (Chang et al., 2005; Franks et al., 1991;
Slade and Levine, 1991). The ‘‘water substitute’’ hypotesis
states that stabilizers can form hydrogen bonds at specific
sites on the surface of the proteins and substitute for the
thermodynamic stabilization function of water that is lost
during drying (Carpenter and Crowe, 1989; Carpenter et al.,
1993; Chang et al., 2005). Following this theory, Strambini
and Gabellieri (1996) proposed that the removal of tightly
bound water from the protein surface by excessive
dehydration correlated with increased rates of protein
aggregate formation during storage. The addition of the
carbohydrate excipients sucrose or trehalose to the
formulation provided a solid-state environment where
complete coverage of protein surface-accessible hydrogen
binding sites was achieved. This correlated with improved
native-like solid-state protein structure and reduced protein
aggregation during storage (Strambini and Gabellieri, 1996).
Strategies for Aggregate Removal and
Minimization During mAb Manufacturing
Processes
During manufacturing, it may not be possible in all cases to
completely prevent or suppress aggregation, so that effective
removal methods are essential for overall protein aggregate
Table I.
management. Numerous strategies are employed to minimize the level of aggregates in bio-therapeutics including
protein engineering, expression system selection, optimization, separation during downstream processing, and storage
formulation buffer screening. Separation during downstream processing is of particular interest as it provides the
biggest opportunity to remove aggregates once they have
been generated. Thus, different purification strategies can be
effectively used to deal with increases in aggregate levels due
to process-driven modifications or other unexpected
changes that may occur during technology transfer and
scale-up. Strategies for removal of protein aggregates need to
be evaluated on a case-by-case basis, however, in this section
we intend to provide a summary of the available strategies to
minimize or reduce aggregates levels across mAbs manufacturing processes.
Chromatography as a Tool to Reduce Aggregates Levels
During downstream manufacturing, chromatography is
typically the step that mostly contributes to aggregate
removals (Table I). The choice of a particular resin and
mode of operation should be guided by fit and compatibility
with the overall process purification train as well as an
appropriate balance of productivity, yield, and product
quality.
Protein A affinity chromatography is typically used as the
first downstream step in purification of therapeutic mAb.
This purification step is not capable of removing aggregates
because product aggregates might bind to the resin ligand as
well as monomer forms of product. Although this step is
unable to remove aggregates, there are several studies
intended to decrease aggregates formation during the
Protein A chromatography step. Sodium chloride (0.1–
1.0 M) has been mentioned as an elution buffer additive to
increase the elution pH (Carpenter et al., 1999; Gagnon,
1995). Ethylene glycol has been used to weaken hydrophobic
interactions and thus increase elution pH from Protein A
columns (Bywater et al., 1983). Another approach is to reengineer the Protein A ligand to allow for milder elution
conditions. This allowed IgGs to be eluted at a pH of 4.5
instead of pH 3.0 (Gulich et al., 2000).
The use of anion- and cation-exchange chromatography
has been demonstrated to be useful at production scale to
Chromatography as a tool to decrease aggregates levels.
Type of chromatography
Objective
How the objective is achieved
Affinity (Protein A)
Minimize aggregates formation
Anion exchange
Cation exchange
Decrease aggregates levels
Decrease aggregates levels
SEC
Hydrophobic interaction
Decrease aggregates levels
Decrease aggregates levels
The use of NaCl or ethylene glycol increase elution pH
Re-engeneering protein A ligand will allow higher elution pH
At protein PI, aggregates bind to resin and product flow-through
Product elutes earlier than aggregates. Only elution peak fraction
containing the product is collected
Monomer has a smaller size than aggregates and elutes earlier
Aggregates are more hydrophobic than monomers and are more strongly
bound to the resin. The use of gradients or more volume of buffer
can be used to first elute the product and later the aggregates
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Biotechnology and Bioengineering, Vol. 108, No. 7, July, 2011
separate mAb monomers from dimers and larger molecular
weight species (Ansaldi and Lester, 2003). When the
working pH is close to the product’s pI (around 0.2 logs
lower than the pI), the overall net charge of the product is
low. Aggregates generally carry more charge than the
product at this pH range and, therefore, if the sample is
loaded on a Q Sepharose column (anion exchange resin) at
these conditions, most of the charged aggregates will be
bound to the column, resulting in removal of aggregates
(Wan and Wang, 2001). A novel class of dextran-grafted
agarose based ion exchangers has gained popularity for
process scale bioseparations. These matrices have been
shown to exhibit both high equilibrium binding capacity
and rapid protein mass transport kinetics. This is of interest
as dextran-grafting dramatically impacts apparent pore
radius as determined using inverse size exclusion chromatography (Harinarayan et al., 2006; Yao and Lenhoff, 2006).
At pH values that are not the pI of the protein, the charge
properties of protein aggregates are expected to be similar to
the monomer but the size will be quite different, it is
reasonable to anticipate that dextran-grafting may influence
the separation of these two forms (Suda et al., 2009).
Aggregates bind more strongly to ion exchangers than the
corresponding monomeric form. For separation of aggregates, similar monomer pool purity and yield could be
achieved with SP Sepharose XL and SP Sepharose FF (Suda
et al., 2009).
Some antibody purification processes use size exclusion
chromatography (SEC) to reduce the levels of aggregates in
the final solution. By employing either Sephacryl 300 or
Superdex 200, gel filtration resins that separate according to
size, it is possible to achieve final product pools with <2%
aggregates (Phillips et al., 2001; Wang et al., 2006). Although
it is technically possible to reduce the level of aggregates in a
protein solution using SEC, it is often not cost efficient to do
so. Preparative SEC is typically inefficient because of the
poor resolution of aggregates from monomer (Litzen et al.,
1993; Wang et al., 2006).
Alternatively, it is feasible to separate mAb from
aggregates based on differences in hydrophobicity, which
has been mainly used for the removal of both aggregates and
impurities such as HCP (Lu et al., 2009). Hydrophobicity of
mAb increases with aggregation, a fact that has significant
theoretical as well as practical significance (Suda et al.,
2009). In the market there are several resins with low to high
hydrophobicity.
Charged-hydrophobic mixed mode chromatography
methods have been applied to antibody purification for
decades and have focused more recently on aggregate
removal capacity. They exploit various combinations of
alkyl and aromatic hydrophobic groups with positively and/
or negatively charged residues (Gagnon, 2009a).
Hydroxyapatite has also been reported to provide good
capabilities for aggregate removal; aggregates levels greater
than 60% have been reduced to less than 0.1% using this
resin (Gagnon, 2009b). Phosphate gradients have also been
proven to remove aggregates in IgG mAb. Effective aggregate
removal is obtained with most antibodies that can be eluted
at 5–15 mM phosphate, but declines in parallel with
removal of HCP, DNA, and endotoxin for antibodies that
require higher phosphate concentrations (Gagnon, 2009b).
Hydroxyapatite chromatography in the presence of polyethylene glycol (PEG) provides a valuable enabling method
for removing aggregates from IgG and IgM mAb (Gagnon,
2008). PEG preferentially enhances aggregate retention,
thereby, increasing the degree of separation between
aggregated and non-aggregated antibodies.
How to Minimize Aggregates Formation During
mAb Processing
Treatments with solvents and detergents had been successfully used to inactivate virus as an alternative to low pH
inactivation. It has been shown that solvents and detergent
treatments of a recombinant protein can completely and
rapidly inactivate enveloped viruses (Horowitz et al., 1998).
The application of these methods instead of the low pH
exposure will reduce the level of aggregates in the product.
Pumping solutions containing mAbs is a common
practice in large-scale manufacturing processes. This fact
implies applying shear stress to mAbs that cause aggregates
formation. The use of lobular pumps is very extended at
large-scale protein purification processes. When a protein
solution is pumped using a lobular pump, the protein is
exposed to shear stress forces ejected by the pump lobes. By
reducing the distance of the pump heads, shear stress will be
reduced and hence aggregates formation (Gomme et al.,
2006a,b).
UF has also been used to remove aggregates in mAb
processes (Wan et al., 2005). A minimal wall shear stress has
to be applied to ensure adequate back transport of deposited
solutes to the retentate but, at the same time, it has to be
limited to avoid the generation of soluble and insoluble
aggregates which adversely affect bulk quality. Rosenberg
et al. (2009) reported that if flow and pressure values are
adjusted depending on the prevailing retentate concentration, the resulting stage process shows an improved
permeate flux and reduced concentration time, also showing
reduced aggregation compared to other methods operating
under constant pressure and flow conditions.
Filtration is a common strategy used to remove insoluble
aggregates. Different studies have shown that aggregates are
precipitated better than monomers by ammonium sulphate,
also indicating that aggregates bind more strongly to PDVF
membranes (Wang et al., 2006).
Agitation has been reported to be a source of aggregates
formation, but some studies have demonstrated that the
magnitude and duration of shear exposure, in the absence of
the air–water interface, does not cause protein aggregation
(Harrison et al., 2003). Therefore, if solutions containing
proteins are agitated gently (avoiding vortex formation),
aggregation will be minimized. Also, it may be possible to
inhibit aggregation induced by agitation by careful choice of
excipients (Serno et al., 2010).
Vázquez-Rey and Lang: Aggregates in mAb Manufacturing Processes
Biotechnology and Bioengineering
1503
mAbs are also exposed to gas–liquid interface when they
are transferred to hold tanks. Typically large-scale mAbs
manufacturing processes use hold tanks to store intermediate process pools. When intermediate pools are being
collected into hold tanks, it is essential to avoid product
splashing, therefore, pool hold tank should have dip tubes or
product inlet pipes designed in such a way that product is
directed toward tank walls. Another solution is to fill tanks
from the bottom instead from the top, in order to avoid
denaturing proteins when filling pool hold tanks. In the UF
systems, it is not recommended to concentrate product to
volumes that are below recirculation retentate dip tube in
order to minimize product foaming and hence aggregates
formation, especially if this step is located downstream
aggregates reduction chromatography.
Surfactants such as the non-ionic surfactants polysorbate
(20 and 80) are the most widely used excipients to prevent
protein aggregation induced by agitation, which has been
explained through different mechanisms. The adsorption
competition between the surfactant and the protein to the
gas–liquid or solid–liquid interfaces has been suggested to
minimize the exposure of protein to these interfaces and,
therefore, protecting from surface induced denaturation and
aggregation (Bam et al., 1998; Carpenter et al., 1999).
Surfactant molecules may interact with the exposed
hydrophobic regions of the protein and, therefore, cover
such sites that cause aggregation (Mahler et al., 2005). The
most likely mechanism for polysorbate inhibition of rhGH
aggregation due to agitation is a steric effect caused by
polysorbate blocking aggregation-prone hydrophobic sites
on the protein surface (Bam et al., 1998). The protective
nature of polysorbate 20 was clearly observed for samples
subjected to shaking stress whilst stirring required much
higher concentrations of polysorbate 20 in order to
demonstrate a protective effect (Kiese et al., 2008).
In order to avoid aggregates formation during freezing, an
ideal strategy is to freeze the entire solution at the same time
and rapidly to minimize thermal transitions (such as
eutectic melting) and glass transitions. With this strategy
solute molecules will be entrapped and prolonged freezeconcentration stress will be prevented (Webb et al., 2002).
The exposure of non-polar surfaces reduces the entropy
and enthalpy of the system, at high as well as at low
temperatures. It is, therefore, important to store biophar-
Table II.
Having discussed aggregate formation and its removal
during bioprocessing the overall goal of any manufacturing
process will always be the achievement of highest possible
product yield and—purity with minimal aggregate levels.
Tables I and II shows a list of recommended strategies that
can be used to minimize or reduce aggregates levels across
mAbs manufacturing process. These data can be used to aid
process development phase and/or equipment selection.
The concept of quality by design (QbD) was introduced in
2004 as a result of the cGMP for the 21st Century Initiative
with the objective of achieving a desired state for
pharmaceutical manufacturing (Rathore, 2009). QbD is
defined in the ICH Q8 guideline as ‘‘a systematic approach
to development that begins with predefined objectives and
emphasizes product and process understanding and process
control, based on sound science and quality risk management’’ (International Conference of Harmonization (2008)
ICH Harmonized
Tripartite Guideline: Q8(R1)
Pharmaceutical Development (http://www.ich.org/LOB/
media/MEDIA4986.pdf)). The main benefits of this QbD
system are to (1) assure product quality through design and
performance-based specifications, (2) facilitate continuous
improvement and reduce chemical manufacturing control
(CMC) supplements, (3) enhance the quality of CMC
reviews through standardized review questions, and (4)
reduce CMC review time when applicants submit a quality
overall summary (QOS) that addresses the questions (Yu,
2007).
How to minimize aggregates formation
Viral inactivation
Pumping
Tank design
Ultrafiltration
Agitation
Freeze
1504
Future Outlook
How to minimize aggregates formation.
Process step
Storage
maceuticals well below their thermal unfolding temperature,
typically at 2–88C in order to minimize aggregates during
storage process (Fesinmeyer et al., 2009).
The appropriate formulation becomes critical when bulk
freezing and thawing is required, and excipients may serve as
protein cryo-protectants (Pikal-Cleland and Carpenter, 2001).
As discussed in Strategies for Aggregate Removal and
Minimization During mAb Manufacturing Processes
Section, many of the mAb are delivered to patients filled
into vials. Brych et al. (2010) showed that eliminating the
headspace prevented the aggregation of IgG A in shaken vials
and this may be a better approach than a label saying ‘‘Do
not shake.’’
Use of solvents and detergents to inactivate virus
Minimize the use of pumps if possible, for example, pressurizing tanks to transfer product
When using lobular pumps, reduce the pump heads distance to minimize shear stress
Avoid product splashing
Adjust flow and pressure values depending on retentate concentration
Avoid vortex formation and or use of surfactants like polysorbate 20 or 80
Use of cryo-protectors
Freeze the entire solution at the same time and rapidly
Eliminate the headspace in vials and avoid shaking
Biotechnology and Bioengineering, Vol. 108, No. 7, July, 2011
Question based review involves three primary components (Rathore and Winkle, 2010): (1) process knowledge
that includes a thorough understanding of process inputs
and their impact on performance, (2) the relationship
between the process and a product’s critical quality
attributes (CQA), and (3) the association between CQA
and a product’s clinical properties. The application of the
QbD during process development phase helps to develop a
suitable process control strategy for those quality attributes
that are considered critical. For aggregate removal, the
application of QbD in the development of the hydrophobic
interaction chromatography has been successfully proved by
Jiang et al. (2010). The manufacturing process of a
biotechnological product, however, is a complex multi-step
process where aggregation can be caused during different
stages of the process. Thus improved process understanding
contributing to aggregation is essential and would allow for
a better design of individual process steps—broad design
space needs to be established—and opportunities for
changes in scale, equipment, etc. without prior FDA
approval. Referring to aggregate removal, it might be
recommended to generate multi-dimensional design spaces
for individual process steps as outlined in Tables I and II and
amended by the design space investigation of upstream
manufacturing conditions which have a strong impact on
product quality, aggregate formation, as well.
Nomenclature
CMC
chemical manufacturing control
CQA
FDA
critical quality attribute
US Food and Drug administration
cGMP
pharmaceutical current good manufacturing practice
HCP
host cell proteins
mAb
monoclonal antibodies
PEG
polyethylene glycol
pI
isoelectric point
PVDF
polyvinylidene fluoride
QbD
QOS
quality by design
quality overall summary
rhGH
recombinant human growth hormone
SEC
size exclusion chromatography
UF
ultrafiltration
VEGF
vascular endothelial growth factor
The authors wish to thank Dr. Francisco Leira and Mr. Ashley
Westlake for their insightful comments and discussion. In addition
they greatly appreciate the critical review of the manuscript by the
journal reviewers. The authors who have been employees by Lonza at
the time of project execution, furthermore, declare no competing
financial interests.
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