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447
Development and Production of Commercial Therapeutic Monoclonal Antibodies in Mammalian Cell Expression Systems: An Overview of the Current Upstream Technologies
Michel Chartrain* and Lily Chu*
Department of Bioprocess R&D, Merck Research Laboratories, P.O. Box 2000, Rahway NJ, 07065, USA
Abstract: This article provides an overview of the upstream technologies used in the industrial production of therapeutic
monoclonal antibodies (mAbs) based on the cultivation of mammalian cells. More specifically, in a first section, after a
short discussion of relevant biochemical characteristics of antibodies, we review the cell lines currently employed in
commercial production and the methods of constructing and isolating production clones. This is followed with a review of
the most current methods of commercial scale production and their associated technologies. Selected references and short
discussions pertaining to emerging and relevant technologies have been embedded throughout the text in order to give a
sense of the overall direction the field is taking.
INTRODUCTION
Therapeutic monoclonal antibodies (mAbs) are as of today a well accepted class of therapeutics especially in the
fields of oncology, immunology, and organ transplant, where
the use of these targeted biologics has profoundly revolutionized treatments paradigms. This is especially true in the
treatment of various cancers where mAbs have proven to
carry fewer side effects than the traditional cytotoxic drugs
and have resulted in improved patient quality of life [1-3].
As of today, there are twenty two therapeutic mAbs, or
fragments, currently registered for marketing in the US (see
table 1) with a market size over $ 17B in 2007 and an expected cumulative annual rate of growth of 14 % [4, 5] . This
success is likely to carry for many years since it has recently
been reported that the number of mAbs that entered clinical
studies tripled over the past decade [4].
achieving high efficiency in these process steps and will
briefly discuss potential technological improvements which
are likely to benefit the current technology over the next few
years.
1. OVERVIEW OF RELEVANT BIOCHEMICAL
FEATURES AND PROPERTIES OF MONOCLONAL
ANTIBODIES
This section will familiarize the reader with the basic
concepts that are later discussed in the following sections.
1.1. The Concept of Therapeutic Antibody
Commercial production of currently registered mAbs
relies on the cultivation of mammalian cells that have been
genetically engineered to over-produce the mAb of interest.
One exception is the use of the bacterium Escherichia coli in
the production of fragments. The production of mAbs at
industrial scale is a multifaceted endeavor that encompasses
many technically complex and lengthy steps. Briefly, a commercial mAb production process starts with the generation of
a mAb via immunization of an animal or via molecular biology methods, the identification and optimization of the coding DNA sequence and the construction and identification of
a highly producing and stable clone. These steps are followed with the development of a well designed cultivation
process that encompasses the full control and scale up of
associated operations that will support early clinical evaluations. Typical cycle times for these activities usually range
between 16-24 months [6]. In this article, we will provide an
overview of the state of the current technologies that lead to
Via interaction with specialized components of the immune system, the in vivo role of antibodies is to clear the
host from invading pathogens and from any non-self molecules these micro-organisms may release (toxins for example). Antibodies present exquisite specificity for their target
(antigens), with the ability to recognize and bind exclusively
to a small region (epitope) of a given antigen. When binding
to an antigen, the complex formed allows for rapid recognition and clearance by specialized components and cells of
the immune system. Natural killer cells NK or NKC recognize antibody-target cell complexes and trigger the lysis and
destruction of the invading cell in a process known as antibody dependent cell-mediated cytotoxicity (ADCC). Another
facet of the immune response involves the complement, a
multi-protein complex that sequentially bind to the antibodytarget complex leading to either its recognition and engulfment by macrophages in a process known as opsonization, or to the lysis of the target cell in a process known as
complement-dependent cell cytotoxicity (CDC). This very
succinct and simplified overview of the immune response
can be supplemented by consulting the following references
for more extensive discussions [7, 8].
*Address correspondence to these authors at the Department of Bioprocess
R&D, Merck Research Laboratories, P.O. Box 2000, Rahway NJ, 07065,
USA; E-mails: [email protected]; [email protected]
These properties have made antibodies a very attractive
choice for novel therapeutic approaches to diseases where
externally exposed membrane-bound or circulating proteins
could be specifically targeted for a specific action. The over-
1389-2010/08 $55.00+.00
© 2008 Bentham Science Publishers Ltd.
448 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
all expectation is that specific interactions between the mAb
and its specific circulating or membrane-bound protein target
will either result in the blockage of key signals of the disease
pathology, will achieve the specific delivery of conjugated
small molecules such as cytotoxins or radionucleides to target cells, or will lead to the destruction of the target cell via
ADCC or CDC. Several therapeutic mAbs whose mode of
action fits one or several of these desired traits are currently
in clinical use. A few are highlighted here.
For example, the mAbs cetuximab (Erbitux®) and trastazumab (Herceptin®) interfere with the over-expressed membrane-bound growth factor receptors EGFR and Her2, both
known to be implicated in signaling for proliferation and
survival of cancer cells. By preventing ligand binding,
cetuximab prevents cancer cells from proliferating. Although
the exact mode of action is still unclear, trastazumab is believed to act via interactions with the effectors of the immune system. The association of the mAb with a membrane
protein triggers various mechanisms that can lead to the destruction of the targeted cell by triggering apoptosis (a preprogrammed self destruction pathway) or cell lysis via CDC
or ADCC. Both these two mAbs are two highly successful
clinical examples of interference with a membrane-bound
receptor. Based on a different approach, rather than directly
interfering with a receptor, the mAbs bevacizumab
(Avastin®) and adalimumab (Humira®) respectively block
tumor angiogenesis and the signals leading to rheumatoid
arthritis by respectively sequestering the circulating ligand,
VEGF and TNF. The concept of delivering a toxic payload
to specific cells targeted for destruction is exemplified by
three mAbs that are currently in use in the treatment of cancers. Mylotarg® (gemtuzumab) is conjugated to ozogamycin, a derivative of the antitumor-antibiotic calicheamycin
and targets CD33, a protein often over-expressed on cancerous myeloid cells. Bexxar® (tositumomab) conjugated with
the radionucleide 131I, and Zevalin® (ibritumomab) conjugated with either 111In or 90Y both target CD20, an antigen
expressed on the surface of myeloid cells. Palivizumab
(Synagis®) which is directed at the envelope transmembrane F (fusion) protein of the respiratory syncytial virus (RSV is a very serious pediatric infection) is the only
mAb currently in use for the treatment of infectious disease.
Fragments of mAbs that are made up of the domain of
the mAb that confers affinity to its target but that do not include the domain of the molecule that interacts with the effectors of the immune system are also used. In addition to
not having interactions with the effectors of the immune system, fragments are rapidly cleared from the body, a second
feature that is useful when seeking short term therapeutic
action. Three fragments are currently registered with the
FDA. Reo-pro® (abciximab) is used to control platelet aggregation during percutaneous coronary intervention. Lucentis® (ranibizumab) which is directly delivered to the eye has
been very successfully used in the management of age related wet macular degeneration. Very recently, the approval
of Cimzia® (certolizumab pegol), a pegylated fragment directed at TNF, marked the introduction of fragments designed for extended in vivo life time through conjugation
with the polymer PEG. These few selected descriptions exemplify how mAbs have been successfully used in the treatment of complex and serious diseases. The list of the 22 cur-
Chartrain and Chu
rently US approved mAbs, their protein target and their
clinical applications can be found in Table 1. Additional
general information can be found in the following publications [9-16].
1.2. Biochemical Overview of Therapeutic Antibodies
As of today all therapeutic mAbs are of the Immunoglobulin G (IgG) sub-class. These molecules are made up of
two heavy chains and two light chains that are held together
and folded via intra and inter-chain disulfide bounds. The
average molecular weight of each sub-chain is about 25Kd
(or about 220 amino acids) for the light chain and 50 Kd
(about 450 amino acids) for the heavy chain. The heavy
chain is made up of 3 constant and one variable domains,
while the light chain is comprised of one constant and one
variable domains. Specificity of the antibody molecule is
dictated by the amino acid sequence at specialized sites,
called complementarity determining regions (CDR) present
in the variable domains of both light and heavy chains. It is
these loop-forming hyper-variable regions of the molecule
that interact and form a tight association with the intended
antigen target. All IgG sub-class antibodies are based on this
general structure and mostly differ in the sequence of the
constant regions of the heavy chains, the part of the molecule
that interacts with other components of the immune system
via ADCC and/or CDC. The IgG1 sub-class molecules present the strongest association with the receptors of the effector components of the immune system [17]. IgG antibodies
have an additional layer of complexity through a conserved
N-linked glycosylation site at ASN-297 on each of the heavy
chains [18]. These short glycans are of bi-antennary structure
and can be variable in many ways that include; the terminal
sugar composition (presence or absence of galactose, capping with a sialic acid) and the presence or absence of bisecting fucose and N-acetyl glucose amine residues. This high
degree of variability and relative abundance of each glycan
species results in the high likelihood that glycans of a different structure are attached to each of the heavy chains of a
given mAb molecule. The presence and composition of these
glycans has been found to greatly affect the activity of the
antibodies in terms of their interactions with the immune
system effector functions [19]. For example, glycans devoid
of the bisecting fucose will confer the mAb a stronger interaction between the antibody and the NKC, resulting in
higher ADCC mediated immune response [20-26]. Similar
increases in ADCC function have also been noticed when the
bisecting N-acetyl glucose amine is present, although this
effect seems to be more antibody dependent [27-29]. However while all ADCC activity is abolished by the complete
removal of the glycans, partial removal of the terminal sialic
acid and galactose have no gross apparent impact on ADCC
[30]. Although they represent a small part of the antibody
molecule, glycans, because they are assembled post translationally are more likely to exhibit higher diversity and to
thereby influence the overall activity of the mAb. Fig. (1)
presents a composite pictorial of an antibody with a composite of the glycan chain presented in detail.
In addition, it is also important to note that not only are
antibody molecules highly complex in their primary structure
and heterogeneity; they also have complex secondary and
tertiary structures that greatly influence their activity. This
Development and Production of Commercial Therapeutic Monoclonal Antibodies
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
449
Fig. (1). Schematic representation of an IgG mAb.
The molecule is symmetric and can engage through its binding regions with two antigens at once. The presence of the hinge region allows
for flexibility and facilitates this process.
The insert provides details of the glycan structure. Symbols key is as follows:
Fucose (FUC
), N acetyl glucose amine (NAGA ) galactose (GAL
), mannose (MAN ), N-acetylneuramic acid or sialic acid
(NANA
).
short overview of the biochemistry of IgG antibodies highlights the fact that synthesis of biologically active mAbs is a
highly complex and well orchestrated biochemical process
that encompasses folding, trafficking, post-translational
modifications, and effective secretion. The choices of a producing platform (host cell) and of the physico-chemical conditions that will support mAb synthesis have been shown to
greatly affect the abundance and quality of the N-linked glycans, molecule folding, and the yield of the mAb of interest.
Since the advent of mAb production in the late 1970s,
production platforms have relied on mammalian cells. As
discussed above, monoclonal antibodies are highly complex
molecules that require post translational modifications such
as inter and intra-chain disulfide bond formations, and the
addition of N-linked glycans at specific sites. Until recent
metabolic engineering advances, only mammalian cells
which possess the intrinsic required machinery could produce mAbs with the desired folding and post-translational
modifications, and were therefore the de facto production
platforms for mAbs. Although considerable progress has
recently been (and continues to be) made in the use of other
biological production platforms [31-33], to our knowledge,
as of today only mammalian cells are used for the production
of commercial and late stage clinical mAb supplies. Judging
by the pace and the significance of the discoveries made in
the use of yeasts, it is however very likely that during the
next decade, these non-mammalian expression systems will
be in use for the production of clinical or commercial mAbs.
2. GENERATION AND MOLECULAR BIOLOGY
BASED MANIPULATIONS OF MABS.
2.1. General Principles
Traditionally, the generation of most therapeutic antibodies relies on the immunization of mice or other mammals
with the desired antigen target. Upon repeated injections, the
animals develop a strong response to the antigen and will
present a large number of cells secreting antibodies against
the injected antigen. It is important to understand that multiple antibodies directed at different epitopes of the antigen are
secreted by a mixed population of B cells with each cell only
secreting one specific antibody [34]. Therefore if one secreting B cell was to be isolated and expanded (cloned), only
one type of antibody molecule (monoclonal or mAb) would
be secreted. Unfortunately, secreting B cells can only replicate a limited number of times, therefore rendering production of mAbs by their cultivation all but impossible. In vitro
production of mAbs was only a concept until the groundbreaking hybridoma technology was developed in the 1970s
by Kohler and Milstein [35-37]. Antibody secreting cells
originating from the spleen of an animal immunized with the
antigen target of choice are fused with immortalized (i.e.
cells that will divide forever when cultivated in permissive
conditions) non-antibody secreting cells. To facilitate the
isolation of the desired fused clones, the immortalized nonsecreting cells are engineered to be deficient in a key metabolic pathway. For example, deficiencies in the hypoxan-
450 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
thine guanidine phosphorybosyl transferase (HGPRT), an
enzyme that holds a key role in the synthesis of guanidine
tri-phosphate (GTP) in the "nucleotide synthesis salvage
pathway" have been extensively used. These cells are still
able to synthesize their nucleotides via the main (or de novo)
pathway, however, when cultivated in the presence of
aminopterin, an inhibitor of dehydrofolate reductase (DHFR)
an enzyme that produces tetrahydrofolate, a key co-factor in
the synthesis of thymidine tri-phosphate (TTP) from dUMP
in the main nucleotide synthesis pathway, these cells can not
grow since each of the pathways leading to the production of
nucleotides are non-functional under these conditions. Since
they have a functioning HPRT enzyme, B cells originating
from the spleen are able to survive in the presence of
aminopterin. However as somatic cells, they only can replicate a few times before dying. Only fusion clones that have
inherited the ability to replicate indefinitely from the nonsecreting cells and the functional HPRT activity from the Bcells will grow for extensive number of generations in the
presence of aminopterin. When coupled with the isolation of
colonies originating from one fused cell, this technology
rendered possible the production of mAbs. Clones that produce the mAbs with affinity to the desired target can be identified using classical affinity biochemistry methods. The development of this technology in the late 1970s was hailed as
a major breakthrough and opened the door to generation of
therapeutic monoclonal antibodies and is now a well established protocol [38-40]. Fig. (2) represents an overview of
the process of mAb production by generating hybridomas.
Although the first recombinant mAbs were produced
using this technology, the mAb hybridoma production platform presents some serious drawbacks. Since the coding
region of the mAb directly originates from the immunized
animal without any subsequent genetic manipulations, the
mAb produced is of mouse origin in its amino acid sequence,
Chartrain and Chu
and upon repeated injections rapidly triggers an immune
response from the human recipient (referred to as Human
Anti Mouse Antibody or HAMA). Upon these repeated infusions, the therapeutic mAb is rapidly cleared from the circulation and therefore rendered therapeutically ineffective.
About 80% of the patients develop an immune response to
the first commercial therapeutic mAb, OKT3, a murine antibody used in the treatment of organ transplant rejection [41].
Further improvements were obviously needed in order to
make monoclonal antibodies a therapeutic reality, and consequently there are only two murine therapeutic monoclonal
antibodies approved at this time (Table 1).
2.2. Sequence Humanization
Following these first clinical attempts to use mAbs of
murine structure, molecular biology manipulations greatly
improved the immunogenic profiles of the subsequent mAbs
tested in humans by reducing the amount of amino acid sequences of mouse origin while retaining the appropriate affinity for the intended target. The most common approach
starts with the generation of mouse hybridomas as described
above. Once clones are generated, they are screened for their
ability to produce a mAb with the desired affinity for the
target antigen, employing biochemical screening methods
[42-47]. The selected clone is further cultivated and the mAb
genetic coding sequences are isolated and modified to yield
antibodies that are more and more human-like in their structure. Briefly these technologies have evolved over the past
decade, from first the construction of chimeric antibodies,
made up of the constant regions of human antibodies attached to the variable region of mouse origin (responsible for
the binding to the target), to techniques of generating "humanized" antibodies by "grafting" and "veneering" only the
essential mouse amino acid residues needed for affinity
Fig. (2). Overview of the various steps leading to the production of mAbs via the generation of hybridomas.
Development and Production of Commercial Therapeutic Monoclonal Antibodies
Table 1.
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
451
Therapeutic Monoclonal Antibodies and Antibody Fragments on the US Market
Name
Generic name
Target
Indication
Rituxan®
Rituximab
CD20, a protein expressed on B cells but not on stem cell and differentiated
antibody secreted cells. Can trigger cell death via ADCC, CDC and by initiating
apoptosis.
Non Hodgkin
Lymphoma of B-Cells
Zevalin®
Ibritumomab
Conjugated with either
Indium 111 or yttrium 90. A
murine antibody precursor
to rituximab
CD20. Same target as rituximab, often used in conjunction with rituximab
Non Hodgkin
Lymphoma of B-Cells
Bexxar®
Tositumomab, a conjugate
with I131
CD20. Specifically delivers a radionucleide to target cells. ADCC, CDC, and
induction of apotosis via ionizing radiation.
Non Hodgkin
Lymphoma of B-Cells
Herceptin®
Trastazumab
Her2, a tyrosine kinase receptor over-expressed on the surface of cancer cells. By
blocking the most upstream signal, it reduces cell proliferation and survival. Can
also trigger cell death via ADCC and CDC.
Cancer (breast)
Mylotarg®
Gemtuzumab ozogamicin
A conjugate with the cytotoxic ozogamycin,
CD33, a protein often over-expressed on cancerous myeloid cells but not on
pluripotent haematopoetic stem cells. Specifically delivers a cytotoxic drug.
Cancer (Acute myeloid leukemia)
Campath®
Alemtuzumab
CD52, a protein expressed on the surface of differentiating lymphocytes. Ellicits
cells death via ADCC, CDC, and the triggering of apoptosis.
Cancer (Chronic
myeloid leukemia)
Erbitux®
Cetuximab
EGFr, a tyrosine kinase receptor over-expressed on the surface of cancer cells.
By blocking the most upstream signal, it reduces cell proliferation and survival.
Can also trigger cell death via ADCC and CDC.
Cancer (Colon)
Vectibix®
Panitimumab
EGFr
Cancer (colon)
Avastin®
Bevacizumab
VEGF, a circulating protein that triggers vascularization, a needed step for tumor
development
Cancer (Colon)
Orthoclone®
OKT 3
Mururomab
CD3,a protein associated with T cell receptors. Steric inhibition by OKT3 induces pan-T cell depletion
Kidney transplant
rejection prevention
Zenapax®
Daclizumab
CD25, an IL2 receptor present on the surface of activated but not resting Tlymphocytes, thereby blocking activation of cytotoxic lymphocytes involved in
transplant rejection
Kidney transplant
rejection prevention
Simulect®
Basiliximab
CD25
Kidney transplant
rejection prevention
Remicade®
Infliximab
TNF, a proinflamotory cytokine involved in various inflammatory chronic diseases
Crohn’s disease, RA
Humira®
Adalimumab
TNF
RA
Xolair®
Omalizumab
IgE, prevent the binding of this proinflamatory signaling immunoglubuling with
its receptor
Allergy related
asthma
Raptiva®
Efalizumab
CD11a, a subunit of the T cell integrin LFA-1, a ICAM receptor involved in Tcell activation and migration, a hallmark of psoriasis.
Psoriasis
Tysabri®
Natalizumab
4 subunit of VLA-4 integrin of activated T cells, preventing interaction with its
brain endothelium counter receptor VCAM-1,thereby preventing T-cells from
crossing the blood brain barrier.
Multiple sclerosis
Synagis®
Palivizumab
F protein (fusion transmembrane protein) of the Respiratory Synsytial Virus.
Binding to the F protein elicits recruitment of the immune system effector components and triggers virus clearance.
RSV infections
Soliris®
Eculizumab
Terminal complement protein C5. Prevents complement mediated lysis of red
blood cells deficient in the regulatory proteins CD55 and
CD59, which makes the cells sensitive to
complement dependent destruction.
Paroxysmal
Nocturnal Haemoglobinuria
452 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
Chartrain and Chu
(Table 1) contd....
Name
Generic name
Target
Indication
Reo-Pro®
Abciximab
(a Fab fragment)
GPIIb/IIIa receptor complex of platelets. Binding to these integrins inhibits platelet aggregation and prevent blood clots formation.
Adjunct in coronary
interventions
Lucentis®
Ranibizumab
(a Fab fragment)
VEGF. Binding with the VEGF ligand prevents interaction with its receptors
VEGFR1 and VEGFR2 on the surface of endothelial cells. When locally injected, It reduces
endothelial cell proliferation, vascular leakage
and the formation of new blood vessels on the central part of the retina.
Wet macular
degenerescence
Cimzia®
Certolizumab pegol
(A Fab fragment conjugated with poly-ethyleneglycol)
TNF-.
Crohn's disease
to the antigen target to a human antibody framework. Finally
the production of fully "human" antibodies through the use
of transgenic mice, which expresses human genes for IgGs,
has been implemented to recently yield its first commercialized mAb, Vectibix® (panitumumab), an IgG2 molecule
[48].
All these approaches which rely on the immunization of
animals require the construction of hybridomas and present
limitations in some situations such as when generating a
mAb directed against a toxin or a highly conserved antigen
across species. To overcome these limitations, the generation
of mAbs by recombinant technology was developed. Post
isolation and amplification of the variable HC and LC genetic coding sequences from B lymphocytes by PCR, it relies
on the construction of highly diverse libraries that have the
potential to generate up to 1010 different mAbs via recombination. Most screening protocols use phage display technology, an approach that by fusing the Fab region coding genes
with the gene of a phage coat protein results in linking genotype to phenotype. Briefly, the phage which displays the
resulting Fab on its coat can be isolated by affinity technologies. After amplification in a bacterium, the genome of the
isolated phage contains the matching DNA sequence which
can be isolated and further manipulated as desired such as
increasing affinity for the antigen target [42-47, 49]. Other
methods such as yeast and bacterial displays have also been
developed and may present a simpler and faster approach to
the development of synthetic mAbs [42-47]. As of today, the
use of phage display technology has been successfully implemented in the generation of fully human antibodies, with
Humira® an antibody directed at the human protein TNF,
being the first to reach commercialization.
Development of the production of chimeric mAbs followed by the refinements described here were the needed
boosts that by greatly reducing or eliminating patient immune response to the therapeutic mAb allowed repeated infusions and propelled therapeutic mAbs to widespread and
successful clinical uses.
2.3. Construction of Genetic Vectors
The construction of producing clones follows the overall
paradigm of classical molecular biology developed for microbial systems. After selection of a hybridoma clone secret-
ing the desired antibody (selection based on affinity for its
target), the DNA sequence coding for the IgG is obtained
and engineered to reduce immunogenicity as described
above. Further refinements aimed at increasing the affinity of
the mAb to its target antigen using phage display technology
can take place at that point during the development cycle of a
mAb [49]. This is sometimes a required step since affinity of
the original murine mAb for its target typically decreases
during the humanization process. The finalized mAb coding
sequences are then inserted on classical genetic vectors
(plasmids) that contain a selectable genetic marker and that
are mass produced using microbial technology. The entire
coding mAb sequence can be enclosed into a single plasmid,
or two plasmids each containing the genes for the heavy
chain (HC) and light chain (LC) respectively can be constructed [50, 51]. The purified plasmid(s) is/are then transfected into non-antibody secreting immortalized mammalian
cells, where upon insertion of one or multiple copies into one
or several chromosomes, the foreign gene is transcribed to
eventually yield secretion of a fully functional mAb. Various
transfection approaches that all rely on the temporal destabilization of the membrane integrity, thus allowing the plasmid
to gain access to the cytoplasm, have been used with success
[52]. Vector design uses the same promotors for both heavy
and light chains and in theory can support similar production
of HC and LC, although more LC may be required since HC
lower transcription efficiency and lower transcript stability
has also been observed. Studies seem to indicate that excess
LC synthesis may lead to better stabilization of the HC and
to higher overall mAb secretion [53]. To the best of our
knowledge, tunable expression systems that favor the transcription and translation of LC or that offer the possibility of
controlling the onset of synthesis via induction have not yet
been commercially implemented although the technology
does exist [54].
Since transfection and integration of genetic material into
the chromosome of mammalian cells are low efficiency
processes, in order to facilitate selection of cells that are
likely to have integrated the foreign DNA and to secrete the
desired mAb, selection protocols are routinely used. These
rely on the principle that when the genes of the mAb and of a
resistance/selectable marker are co-localized on the vector
they will co-integrate and co-express with a high probability.
Usually resistance to an antibiotic or the ability to grow in a
Development and Production of Commercial Therapeutic Monoclonal Antibodies
nutritionally selective medium conferred by a gene cointegrated on the plasmid is used at this step. These strategies
will be specifically discussed in detail along with the description of the cell lines for which they have been implemented.
It has been established that the site of integration of the
foreign DNA clearly influences the level of expression. Targeting highly transcriptionally active regions of the genome
for the integration of the foreign genes has been attempted
and has shown some positive but limited signals [55]. One of
these approaches relies on first developing and isolating high
producing clones where the inserted transgene is flanked
with bacteriophage-derived short DNA strands (lox P) that
are substrate to a specific recombinase (Cre) thereby creating
pre-characterized sites for the future exchange and insertion
of other transgenes. Assuming that integration of any foreign
trans gene to this specific site will yield high producing
clones, targeted integration can be achieved via reciprocal
site specific integration [56]. Other approaches have, rather
than targeting transcriptionally active regions, developed the
use of short nucleotide sequences that help the foreign DNA
to be more transcriptionally active regardless of its site of
integration. When linked to the trans genes these genetic
elements are believed to influence the accessibility of the
chromatin environment in the vicinity of the integration site
and to thereby increase transcription efficiency. Several
technologies have been developed and although different in
their respective mode of action, they all rely on this general
philosophy.
The STAR elements (Stabilizing and Antirepressor elements) reduce the extent of histone deacetylation pattern and
the spread of methylation in the vicinity of the inserted trans
genes. Correlatively, the region is kept acylated and transcriptionally more active [57]. The S/MAR elements (Scaffold/Matrix Associated Regions) interact with the nuclear
matrix and create loops where gene expression is coordinated and insulated from repression [58, 59], while the
UCOE elements (Ubiquitous Chromatin Opening Elements)
which are derived from housekeeping genes, a group of
genes that present high histone acetylation, create a highly
transcriptionally active environment [60].
One very apparent impact of these new technologies is
the potential to reduce the need for the evaluation of a large
number of transfectants which could significantly shorten
development timelines [55]. As of now, although several
publications outline the benefits of these elements, their introduction is too recent to know when they eventually will be
incorporated into the design of a commercially used clone.
Table 2.
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453
3. CURRENT CELL LINES IN USE
Although many immortalized mammalian cells lines can
be potentially used, since they were in use at the time the
first mAbs were developed for clinical and commercial applications, as of today, with the exception of the two mAbs
produced by hybridoma technology, the production of all
currently commercialized therapeutic mAbs is achieved in
mammalian cells of mouse (NS0 and SP2/0 cell lines) or
Chinese hamster ovary (CHO cell line) origin. For references, Table 2 lists a few selected commercialized mAbs and
their respective production platform.
3.1. NS0 and SP2/0 Cell Lines
These cell lines originate from mouse plasmacytoma
cells that have undergone several steps of cloning and selection to yield immortalized non-IgG secreting B cells [36, 37,
61-65]. Both NS0 and SP2/0 cell lines have been extensively
used for the production of mAbs employing fusion technology. More recently, molecular biology techniques have allowed NS0 and SP2/0 cells to be used as the production platform for several therapeutic mAbs currently on the market.
These cells can be cultivated in serum and non-serum containing cultivation media and are reasonably amenable to
scale up in large cultivation vessels (up 20,000-L scale).
However, NS0 cells present some drawbacks that complicate
their cultivation. Unlike most mammalian cell lines, NS0
cells seldom grow in the absence of exogenous cholesterol,
and are therefore routinely cultivated in the presence of cholesterol, which is usually delivered via the use of serum [6670]. Since the presence of ingredients of animal origin as
well as any proteins in cultivation media is undesired, it
complicates the use of NS0 cells due to the difficulty of supplying cholesterol in a protein free medium as it requires the
use of carriers such as cyclodextrins to enhance cholesterol
"solubility" [71, 72]. It has been demonstrated that cholesterol independent NS0 cells can be developed, although with
a certain unpredictability, a factor that has limited their use
[73-75]. Recent advances into the mechanism of cholesterol
requirement has assigned the epigenetic gene silencing
caused by methylation upstream of the region coding for the
the17-hydroxysteroid dehydrogenase type 7 that catalyzes
the conversion of lanosterol to lathosterol as the cause of this
deficiency [76-78]. Although industrial groups have developed cholesterol independent NS0 lineage [79], to the best of
our knowledge, no commercial production of mAbs using
NS0 cholesterol independent cells is currently on-going. It is
quite possible, since they offer greater simplicity of use, that
these biochemically and genetically understood cholesterol
independent lines will eventually be used for commercial
production.
Commercialized mAbs Production Cell Platforms
CHO
NS0
Sp2/0
Murine Hybridomas
Avastin, Campath, Herceptin, Humira, Raptiva, Rituxan,
Vectivbix, Xolair, Zevalin
Mylotarg, Soliris, Synagis, Tysabri,
Zenapax
Erbitux, Remicade,
Reopro, Simulect,
Bexaar, Othoclone
454 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
Chartrain and Chu
Although reasonably similar, the glycosylation pattern
distribution ratio (G0F, G1F, and G2F) of the IgGs produced
by NS0 and SP2/0 is not that of circulating human IgGs. In
addition, these cells produce small amounts of murine-like
glycans such as the addition of an extra galactose (-Gal) to
the terminal galactose and the insertion of N-glycolyneuraminic acid (NGNA) in place of NANA, [80], which
have the potential to trigger an immune response. Clinical
adverse events, anaphylactic shock, have been reported for
cetuximab which is produced by cultivation of SP2/0 cells
[81]. Nevertheless, and because the consistency of the glycoforms can be controlled and characterized, NS0 and SP2/0
cells have demonstrated reliability and have proven their
worth in the production of several well marketed mAbs.
which inhibits the de novo nucleotide synthesis pathway,
only those cells that have incorporated the XGPRT gene will
grow. Since the gene coding the IgG of interest is co-located
with the XGPRT gene on the plasmid, the probability of a
co-integration is high, and transfectants expressing high levels of IgG are likely to be selected by using this method [79,
83].
Several selection strategies for selecting clones that have
integrated the desired mAb trans genes have been developed
and successfully applied. The overall philosophy is to use
cells with a deficient metabolic background, while the genetic element containing the mAb coding region carries a
gene that once expressed will complement the metabolic
deficiency. Upon transfection, one or several copies of the
genes carried on the plasmid will integrate into one or several chromosomes of the recipient cells where they will be
transcribed and translated. Post transfection and during cloning of the cells, a selective pressure aimed at favoring those
cells that have incorporated the gene coding for the correction of the metabolic deficiency is applied in order to only
favor the growth of those cells that have integrated the foreign DNA. Assuming that both foreign genes have stayed
linked during their integration into the genome of the recipient cells, the majority of the cells surviving the selective
environment will be mAb producers. Several variations on
this strategic theme are described bellow.
In the mid-1980s CHO cells were modified to be deficient in the dehydrotetrafolate enzyme (DHFR). This enzyme is required for the synthesis of reduced tetrahydrofolate, a cofactor required in the synthesis of DNA precursors.
Commercial production cell lines are derived from two major CHO dhfr minus lineages, DG44 and DUKX-B11. Both
lines were developed at different times by the same group at
Columbia University [88-90]. This metabolic deficiency
allows for a selection strategy similar in its philosophy as
described previously. Here, the use of metotrexate (MTX),
an inhibitor of DHFR is implemented. Since the plasmid
contains a copy of the DHFR gene along with the mAb of
interest, only the cells that have stably integrated both genes
carried on the plasmid will be able to grow and produce mAb
in the presence of MTX [91]. Using increasing concentrations of MTX will lead to the selection of high producing
clones which are likely to have incorporated high copy numbers of the trans genes [51, 92]. The DHFR selection strategy can be enhanced with the use of resistance to aminogalactoside antibiotics such as neomycin and kanamycin conferred by an added resistance gene on the plasmid [93, 94]. It
is believed that the incorporation of high copy numbers of
the trans gene may lead to genetic instability.
A popular NS0 cell lineage is the GS-NS0 commercialized by Lonza biologics. NS0 cells have very low levels of
the endogenous enzyme glutamine synthetase (GS), and require exogenous glutamine in order to grow. It is only when
transfected with a copy of the GS gene that the cells can
grow in the absence of glutamine [61, 82]. A vector, designed to carry both the genes coding for the mAb to be expressed and for GS, once integrated into the genome will
allow the expression of both genes and the selection of producing clones. In order to impose selective pressure against
rejection of the foreign DNA and to select for clones having
integrated the genes in a region of high expression, the cells
are cultivated in a medium devoid of glutamine and in the
presence of methionine sulfoximide (MSX), an inhibitor of
GS. Only those clones that produce large amounts of GS will
survive. Correlatively, these clones are likely to also produce
large amounts of the mAb. Interestingly this approach seems
to yield high producing clones that have integrated low copynumber (less than five) of the trans gene [61, 82].
Another method of selection used relies on the fact that
SP2/0 and NS0 cells are deficient in HGPRT a negative genetic trait can be exploited by inserting a copy of a microbial
gene (from E. coli) coding for the synthesis of XGPRT, an
enzyme that can substitute for HGPRT. Only the cells that
have integrated the XGPRT gene will be able to use the nucleotide synthesis salvage pathway. When cultivating the
transfectants in a selective medium containing aminopterin
3.2. CHO Cells
These cells originally isolated from a Chinese Hamster
Ovary in 1957, are proline auxotrophs and have been spontaneously immortalized during their successive transfers.
[84-87].
More recently, a selection based on the fact that the parent CHO-K1 cells have a low glutamine synthetase (GS)
expression level has been developed. Transforming the cells
with a plasmid co-expressing GS and an IgG of interest and
cultivating the transfected cells in the absence of glutamine
and in the presence of MSX allows for only those cells that
have stably incorporated the foreign genes to grow. Since it
only relies on a weak expression rather than a metabolic deficiency, this system requires that the presence of MSX be
maintained during cell expansion in order to keep a sufficient
genetic pressure to prevent deletion of the foreign DNA [82].
These two genetic strategies rapidly allowed the use of
CHO cells since it conferred a rapid method to select for
producing clones and to identify high mAb producers. CHO
cells can be cultivated in suspension in serum-free chemically defined cultivation media in large scale conventional
bioreactors. They display a high resilience to cultivation
conditions, do not require cholesterol, and tend to remain
viable for a longer period of time when compared to NS0
cells. This ease of genetic selection coupled with their very
amiable character to large scale industrial cultivation have
resulted in the CHO cell line being used in more than half of
the registered mAb production processes.
Development and Production of Commercial Therapeutic Monoclonal Antibodies
However, because of their rodent origin, the glycosylation pattern distribution ratio (G0F, G1F, G2F) of mAbs
produced by CHO, NS0, and SP2/0 cells do not completely
match that of circulating human IgG1. In addition, CHO
cells produce small amounts of non-human like glycan patterns, such as 2-3 linked sialic acid residues that have the
potential to be immunogenic [80]. On the other hand, these
non-human glycans are present in very low proportions (a
few %) and mAbs produced by cultivation of CHO cells
have remarkably safe profiles in the clinic [41].
3.3. Emerging Mammalian Cell Lines
Other cell lines of animal and non-animal origin have
been considered for expression of mAbs. Since this discussion is limited to mammalian cell lines in its scope, and although some significant advances have been made in using
microbial expression systems, this topic will not be reviewed
here. Among emerging mammalian cell lines, the Per.C6®
cell line appears to be most advanced in its usage and acceptance. Per.C6® cells are human embryonic retina cells that
were immortalized by the use of the early gene E1 of Adenovirus [95]. Per.C6® cells offer the potential for human like
glycosylation pattern with the added advantage of a lack of
undesired murine glycans. Several recent communications
have disclosed that Per.C6® cells can be cultivated to very
high densities at large scale and that they are capable of supporting elevated recombinant protein yields of up to 10 g/L
of a test mAb [96, 97]. These promising data will certainly
help the Per.C6® cell line to gain appeal in the next few years
as a potential platform for the production of recombinant
mAbs. However, to the best of our knowledge, no mAb production platform based on the cultivation of Per.C6® has yet
reached the regulatory registration stage.
A few additional cell lines are believed to be in development such as the avian EBx and the rat myeloma YB2/0 cell
lines. Their potential advantage is significantly reduced fucosylation, and therefore the potential to increase the level of
ADCC of the manufactured antibody. Their development in
the production of mAbs is less advanced than that of PerC6
cell. Unlike well established cell lines such as CHO, NS0,
and SP2/0 that have been used for the production of several
commercial mAbs, new and previously unregistered cell
lines are likely to face high regulatory scrutiny. This is especially true on the topic of associated viruses, and in the
clearance of residual genetic material. We are however positive on the prospect of the most advanced of these additional
cell lines, Per.C6® cells, to soon join NS0, SP2/0 and CHO
cells as platforms for the commercial production of mAbs.
4. SELECTION AND DEVELOPMENT OF HIGH
PRODUCING AND INDUSTRIALLY AMENABLE
CLONES
4.1. Clonality and Selection of High Producing Clones
Once cultivated under selective conditions, the cells are
rapidly subjected to cloning by the use of various methods.
Cloning is required for ensuring that the produced mAb
molecules will have high homogeneity and that cultivation of
the selected clone will be consistent and predictable. In addition to being a process need, clonality is also a regulatory
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
455
requirement. The most popular and simpler method is the use
of limiting dilution where the cells are diluted to concentrations that will likely dispense one or less than one cell per
culture well. The cells are allowed to grow under the desired
selective conditions until confluence is observed. This is
achieved by classical dilutions and estimations of cell population based on viable cell counts. An improvement to this
method is the use of automated cell sorting equipment (flow
cytometer). Although these methods are simple and predictable, they require extensive amounts of labor and time and
improvements or alternative methods have been sought.
Adding to the advantage of simple cell sorting, several technologies have evolved with the aim to directly identify potential high producing cells at this stage. One approach has
been to estimate the secretion potential of a cell by entrapping it in a gel. After an incubation period, the secreted entrapped mAb is detected using a fluorescent-labeled antibody. The entire gel drop, containing cells and their respective captured secreted product, are sorted by the flow cytometer according to their surrogate level of mAb production, thereby potentially achieving a rapid selection [98-100].
Other approaches have relied on detecting the intra-cellular
co-expression of DHFR [101], when employing CHO cells
and this specific popular selection system, or on measuring
an easily detectable mAb co-expressed protein directed to
the external surface of the membrane [102]. However, these
methods only reflect the secretory potential of a cell for a
short amount of time under conditions that may not be highly
reflective of the intended production environment.
In an attempt to potentially better capture the production
potential, other methods rely on the cultivation of the cells
on a soft agar surface, which retains the antibody secreted in
the vicinity of the cells, and on detecting extended production potential using a labeled antibody [103, 104]. The clones
can then be directly picked and cultivated for further evaluation. The potential advantage of this technique is that unlike
the FACS based methods, which submit the cells to high
shear stress, it offers a more cell-friendly environment and
captures antibody secretion over a longer period of time,
thereby increasing accuracy of detection.
In order to verify clonality, microscopic observations of
each culture well are performed during the incubation period
which can last several weeks. The cells are further expanded
in larger cultivation vessels where their potential to produce
large amounts of mAb is evaluated [105-107]. Upon confirmation of their mAb production level and quality (i.e., glycan distribution, correct amino acid sequence, ..) , the best
producing clones are selected for laboratory scale cultivation
under process conditions [108]. Since several hundred clones
are evaluated during a typical screen, as mentioned in the
previous section, the use of genetic elements that can target
the foreign genes to highly transcriptionally active regions
has the potential to tremendously reduce both the amount of
work and the development timelines. Of these clones that
maintain high productivity and that produce mAbs with the
desired quality, a few are selected for further re-cloning. This
is achieved by either a second round of limiting dilution or
by sorting the cells using a flow cytometer or any other
method described above.
456 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
Post second round of cloning, an expansion protocol
similar to the one described above is put in place and the best
producing clones (usually 3-5) are selected for stability
evaluation. Since mammalian cells have a very plastic genome and tend to reject, modify or relocate integrated foreign DNA, it is important to ensure that the clone selected
will exhibit genetic stability over the desired length of cultivation. Stability is not only measured in terms of production
levels but also on the biochemical attributes of the mAb such
as sequence integrity and fidelity in the glycosylation pattern
[109]. This is achieved by cultivating the cells for extended
numbers of generations under conditions that mimic process
conditions as closely as possible, including an evaluation
beyond the number of generations that are targeted in the
manufacturing process. It is not uncommon to observe a reduction in the amount of mAb produced over these successive transfers as well as to detect changes in some of the biochemical characteristics of the molecule [107, 110-114].
Most important to the selection of a clone is maintaining
biochemical profile of the molecule as quality attributes are
essential to the safety and potency of the product. Once these
steps are completed, the selected clone is preserved in a set
of cell banks that upon qualification (purity, lack of adventitious agents, etc.) will be used for commercial production of
the therapeutic mAb.
These procedures are lengthy, require a highly specialized and competent workforce, expensive equipment and
therefore weigh heavily on the overall product development
cycle time, since these activities are performed on the critical
path. This is where the advantage of integrating novel technologies can be of great impact since they have a high probability of reducing the extent and duration of the screening
activities. Fig. (2) presents a pictorial overview of the steps
associated with clone selection.
4.2. Cell Line Engineering
In addition to selecting suitable and stable clones for
commercial production of mAbs, many research groups have
attempted to modify the producing host cells in order to either improve or control the quantity and quality of the mAbs
produced. Rather than presenting an exhaustive list, a few
key examples pertaining to each area of investigation will be
discussed.
Improving or influencing the quality of the monoclonal
produced is of importance since it can translate into better
therapeutic efficiency. As mentioned earlier, of significance
is the importance of the glycosylation pattern. For example,
mAbs with non-fucosylated glycans have been shown to
have greater interaction with the effector cells of the immune
system, translating into greater cell target lysis by ADCC.
CHO cell lines that are devoid of fucosyl transferase have
been constructed by several groups, with the resulting mAbs
exhibiting up to 100 fold increase in ADCC activity [20-26].
Other approaches have targeted the modification of the Fc
region of the protein sequence in order to increase affinity
for the receptor of the effector cells or to improve mAb stability [45, 115, 116]. Since high ADCC activity seems to
correlate with the efficacy and response rate of several mAbs
used in the treatment of cancer especially in the case of Rituxan® [117], there is a reasonable likelihood that one or
Chartrain and Chu
several engineered mAbs with increased effector function
achieved via either glyco or protein engineering will reach
the clinics and the market in the next decade.
In addition to improving the end-product, engineering of
the production host may provide a means to improve mAb
production yields. Investigated areas cover a wide range, and
the following selected examples will briefly present an overview of the current research.
Improving cell longevity should result in increased mAb
production since the specific rate of mAb secretion (mass per
cell per unit of time) tends to be fairly constant during the
cultivation stage. Although cells are adapted to grow in suspension the combination of stressful shear bioreactor environment, the potential nutrient limitations, toxic by-product
accumulation, and the burden of producing large amounts of
a foreign protein all have the potential to trigger the onset of
apoptosis, a highly regulated pre-programmed self destruction process that cells will initiate when facing highly stressful conditions [118, 119]. Since the apoptotic pathway is
highly complex and involves many activator as well as repressor proteins, multiple targets with the aim to reduce cell
death have been evaluated. Mutants that contain additional
copies of genes coding for resistance to apoptosis have been
constructed for both NS0 and CHO cell lines, while other
approaches have relied on the use of inhibitors of apoptosis
[120-127]. Overall increases in longevity have been observed and some correlations with increased mAb secretion
have been observed. These published works, however, have
translated to limited improvements from low production levels to modest production levels at best and remain to be
translated to industrial cell lines. Possible explanations include the fact that since industrial cell lines undergo an extensive, lengthy, and stressful selection process, only the
most resilient cells survive and end up selected, and limited
benefit is gained from these additional alterations. It is
highly possible that in the coming years the use of naïve host
cells engineered to exhibit high resistance to apoptosis will
be used in commercial mAb production. When coupled with
the use of genetic elements and of rapid identification methods for high producers, the introduction of engineered cells
may help in speeding up the time required for the identification of clones that are amenable to cultivation conditions.
Another aspect of cell line engineering currently investigated is the modification of the protein assembly and secretion machinery of the cells. Once foreign genetic material is
integrated into a highly transcriptionally active region, supported by retrospective investigations employing proteomic
and genomic analyses, it is now believed that the assembly
and secretion step may in itself be more rate limiting than the
abundance of mRNA [128, 129]. This hypothesis is supported by the fact that maximum secretion rates for mAbs
have been relatively flat over the past 6-8 years, with most
specific production rates of industrial cell lines comprised in
the 25-40 pg/cell/day range. It is speculated that mis-folded
proteins that are degraded in situ can account for substantial
mAb losses, and it is believed that these yield losses could be
reduced through achieving a better control of the
oxido/reductive enzymatic cell machinery and of the interactions with the chaperone proteins involved in cellular trafficking and secretion of the mAb [130-132]. These very
Development and Production of Commercial Therapeutic Monoclonal Antibodies
novel approaches have yet to reach a technological level
where they can be implemented in the design of improved
industrial cell lines. However, they present long term potential and merit to be closely monitored for their eventual implementation.
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
457
liter, even at industrial scale. This is one of several costs of
goods (development time, cultivation medium, purification
resins, and facilities) that significantly contribute to the overall pricing of therapeutic mAbs.
5.2. Cell Banks and Seed Train Processes
5. CULTIVATION PROCESS
5.1. Cultivation Medium
Media used for the cultivation of mammalian cells are
highly complex and contain several dozens of ingredients
that range from amino acids, vitamins, trace elements, and
nucleosides to name a few groups [133-136]. Development
and manufacturing of these finely balanced media are complex and time consuming and require considerable expertise.
Until recently, the use of serum was standard and provided
the cells with many necessary nutrients and growth supporting molecules. However, recent events linked to abnormal
prions which cause bovine spongiform encephalitis (BSE)
have generated an incentive for the avoidance of animalsourced ingredients in the manufacture of biopharmaceuticals such as therapeutic mAbs [137]. Early serum-free cultivation media used the addition of complex plant or yeast
hydrolysates to their formulation in order to supply the nutrients and growth supporting molecules present in the serum
[138-143]. While these components did replace animalsourced materials, hydrolysates are still relatively undefined
raw materials that can vary significantly between lots and
vendors. Recently, the development of protein-free chemically defined media has been successfully achieved for both
CHO and NS0 cell lines, with the formulation of media supporting extremely high cell densities [144]. Particularly, the
development of non-animal sourced cholesterol carriers such
as methyl--cyclodextrin (mCD) complexed with synthetic
cholesterol have met with success in supporting the cultivation of NS0 cells in serum-free media [71, 73].
Some companies choose to develop their own proprietary
medium for many reasons including cost, control of the medium production process, ability to design medium to fit the
specific clones and to avoid reliance on a single vendor since
the formulations of commercially distributed cultivation media are not in the public domain. In house medium development is a significant investment in time and resources and
often includes not only the design of a basal medium, but
also the formulation of the seed expansion and production
medium as well as the related feed solutions if a fed-batch
strategy is to be implemented . One must also consider if
liquid formulations are sufficient or if powder formulations
should be developed. Given the anticipated scale of manufacturing batches for therapeutic mAbs, the storage of several hundreds of thousands of liters of cultivation medium
under refrigerated conditions is a logistically complex and
expensive proposition. Although not discussed in the public
domain, the development of powder media is highly complex
in itself, with the addition sequence of the ingredients and
the type of milling equipment used believed to be keys to a
successful formulation. When considering the number of
high purity ingredients, the complexity of the manufacturing,
and the required tests implemented for quality and consistency, it should not be surprising that the cost of mammalian
cell cultivation media can reach costs in excess of $20 per
Cells destined for cryopreservation are usually cultivated
in small vessels, harvested during exponential growth phase,
concentrated, and mixed with a cryo-protectant such as
DMSO. Cell banks are traditionally stored using glass or
plastic cryovials that range from 1-5 mL in volume. Very
often, two tiered cell banks are created (Master Cell Bank
and Working Cell Bank) in order to ensure long term supply
for marketed products. The Master Cell Banks are made
large enough to ensure supplies for the entire expected lifetime of the commercial process. Storage logistics and protection of the Master Cell Banks are keys to ensuring consistency of operations and should ensure legacy of the product.
Recent publications have discussed larger volume cell banks
using cryobags (up to 100 mL) [145-147]. Since these start
from a larger volume, such an approach reduces the number
of expansion steps needed in the seed train, which also results in shorter timelines. However, this is still a relatively
new proposal for cell bank generation. A few hurdles remain
regarding industrial acceptance of larger volume cell banks the increased storage space requirements and more importantly maintaining integrity of the bag-containers under cryconditions.
The development of the seed or expansion train is often
overlooked or minimized in its importance. While the seed
train has a simple goal – generation of enough cell mass to
inoculate the final production reactor – there are several important decisions that need to be made during seed train development. These decisions include the following: will the
seed train be regenerated with each production batch, how
many expansion steps will be used, will disposable vessels
be used, what type and size of cell bank will be generated?
There are two common approaches for seed train processes – the rolling seed train and a de novo seed train for
each production batch. Of importance is that mammalian
cells are slow to divide with an average doubling time of
about one day, and must not be excessively diluted when
transferred to a new cultivation vessel. It is usual to consider
a 10 fold dilution factor as maximum and safe for these
transfers, and this will minimize the number of vessel transfers. When taken together, these limitations render the expansion of mammalian cells a very lengthy and labor intensive exercise. Some processes are designed at lower dilution
factors, which allow for more flexibility in inoculation volumes and may shorten the seed train process. A de novo seed
train consists of the thawing of a vial (or several vials) of a
cell bank for each production batch. It is not unusual for de
novo expansion protocols to span over more than one month
when targeting the inoculation of very large bioreactors
(10,000-L or greater). In addition to their length, as their
name indicates, de novo expansion protocols must be implemented for each batch, consume a large number of banked
cell vials over time, can be logistically difficult to manage,
and consume a large amount of resources.
458 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
A "rolling seed train" is generated from cells contained in
a vial (or several vials) that are thawed and sequentially expanded to yield a desired volume that is used to inoculate a
bioreactor. Instead of being operated as a batch, the reactor is
maintained in operation for many months. Every few days, a
substantial volume is drained from the reactor and used to
seed the production bioreactor. An equal volume of fresh
medium is added to the seed reactor, thus allowing the remaining cells to multiply again. This drain-fill protocol can
be repeated every few days, and constitute the basis of the
"rolling seed train". The advantage of a rolling seed train is
that smaller cell banks can be generated since each vial can
support multiple production batches over periods of several
months. In addition, less time is required to generate enough
cell mass for production batches once the rolling seed train is
established. However, in order to use a rolling seed train
successfully, cell lines must be stable over much longer time
periods – i.e. 3-6 months rather than 1-2 months. As discussed previously, the selection of a suitable clone for industrial production of mAbs must take into account the potential
implementation of a rolling seed train. This is a very important point to consider since the quality of the mAb generated
using cells produced by a rolling seed train must be consistent. Additional considerations for implementing a rolling
seed train require that the facility be equipped with additional reactors dedicated to the seed train. On the other hand,
a de novo seed train can be implemented with more flexibility as a facility can use disposable reactors for the seed train.
The number of expansion steps is a function of cell bank
vial size, volume of available bioreactors, cell growth rates,
and desired split ratios. Traditional expansion vessels include
spinner flasks, shake flasks, and stirred tank reactors. Recently, many processes have implemented the incorporation
Chartrain and Chu
of disposable vessels, such as Wave bioreactors©, in order to
build in more flexibility into the expansion process and to
control production costs (disposable vessels will be addressed in Section 5.3.3). A schematic overview of cell bank
preparation protocols and of seed train expansion strategies
is presented in Fig. (3).
5.3. Production Process
5.3.1. Environnemental Conditions
Bioreactor controls are one of the most well-defined areas of mammalian cell culture, encompassing temperature,
pH, O2, CO2, and agitation controls. Initial work with bioreactor control approaches was performed in hybridoma platforms, but these approaches have also been implemented
with success in CHO, NS0 and Sp2/0 platforms. Mammalian
cells are typically cultivated at 37°C, pH 7.15 while maintaining dissolved O2 levels at 30-60% dissolved oxygen
(DO). The accumulation of byproducts can be detrimental to
a culture, particularly when the goal is to maintain the culture over multiple weeks. Typical manufacturing processes
are 12-14 days in length with no design for byproduct removal, so the accumulation of byproducts such as lactate and
ammonium can lead to stressful conditions and higher probability of apoptotic cultures. Therefore, many different approaches have been investigated to reduce byproduct accumulation. One of the benefits of the GS system (see section
3.1) is the elimination of ammonium accumulation by eliminating the need for exogenous addition of glutamine in the
culture medium. Both pH shifts and temperature shifts reduce cellular metabolism which leads to lower lactate production and sometimes even a shift in the lactate accumulation profile. Alternative medium components can also help in
Fig. (3). Cell expansion logistics.
Top part, production of Master and Working cell banks (MCB and WCB).
Bottom part, classical cell expansion protocol and rolling seed strategy.
Development and Production of Commercial Therapeutic Monoclonal Antibodies
controlling byproduct accumulation as many processes have
demonstrated through the substitution of galactose for glucose (see section 5.3.2). Many studies have been performed
to examine the effects of ammonium and lactate on CHO and
NS0 cultures. Some studies show minimal negative effects
on cell growth, but productivity may actually benefit from
increasing lactate [148, 149]. Dissolved CO2 (measured as
partial pressure; pCO2 ) levels are closely monitored due to a
desire to mimic physiological levels (pCO2 = 31-54 mmHg),
however, levels can reach >150 mmHg in high cell density
cultures [150, 151].
In order to maximize mAb production, many bioreactor
processes have been designed with a two-phase approach
where conditions are optimized first for growth and then
altered to optimize mAb production. Unlike microbial cultures, mammalian cells do not typically carry inducible promotors; therefore, these processes cannot be induced in the
traditional sense. On the other hand, bioreactor controls such
as temperature and pH can be used indirectly to control mAb
production either by increasing specific productivity or by
shifting cell resources away from cell growth and towards
mAb production. By designing for a two-phase process, after
the desired cell mass has been accumulated, cell mass is
maintained or incrementally increased, but at a very substantially reduced growth rate. This allows the cells to redirect
the cellular metabolism toward mAb production. Low temperature and low pH are particularly effective in CHO and
hybridoma systems, where temperatures as low as 30-35°C
and pH as low as 6.7-7.0 can reduce cell growth and sometimes even increase specific productivity. Studies have found
that growing CHO cells at lower temperatures may keep
cells in the G1 stage of the cell cycle, which allows for more
protein production, less nutrient consumption, less waste
production (e.g. lactate), and decreased cell death [152-157].
The control of pH is also a critical parameter in controlling
cell metabolism. Through lowering the pH of the cultivation
medium, it is possible to decrease nutrient (especially glucose) consumption rates which in turn greatly reduce waste
(lactate and ammonia) production. Lower pH limits are cell
line and clone dependent where too low a pH will result in
lack of cell growth or even cell death. Typical lower pH
ranges where CHO cells can still proliferate and produce
mAb is 6.7-7.0. The final effect is maintenance of higher
viability cultures over longer periods of time. Some clones
even show increased specific productivity when cultivated at
lower temperatures and lower pH. Since clones vary in their
sensitivity to pH and temperature variations, experiments
must be conducted to balance cell growth with specific productivity as a function of these parameters. In addition, as
discussed in the clone selection section, the identification of
clones fully amenable to industrial cultivation conditions
should ideally include the potential implementation of pH
and/or temperature shifts.
CHO cultures do not show much sensitivity to DO levels
– in most cases, CHO cells can be cultivated at DO levels
ranging from as low as 5% to >60% DO without negative
effects. Generally, O2 levels are maintained through the
sparging of a combination of air and O2 supply and by increasing flow rate and agitation, as oxygen demand increases
during exponential growth. Oxygen uptake rates (OUR) can
be a useful tool for culture monitoring as the OUR values are
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
459
very sensitive to culture conditions and closely reflect cellular metabolism. In fact, control automation programs can be
designed to trigger events based on OUR values.
High cell density cultures can reach high pCO2 levels
(>150 mmHg), and these levels will alter cell growth, protein
production, and protein glycosylation [151, 158]. Some of
these effects are believed to be due to the increase in osmolality resulting from typical pH control strategies. When employing the most commonly used buffer system, bicarbonatebuffered medium, increasing pCO2 resulting from high respiratory cellular activity will result in acidification of the medium. In order to keep the pH value close to the desired neutral range, base addition is required and will result in higher
osmolality. To avoid this undesired cycle, the use of bicarbonate-free buffers were successfully implemented in a perfusion process resulting in 70% reduction in pCO2 levels and
subsequent positive effects on cell growth and specific productivity [159]. However, the negative effects of elevated
pCO2 still appear to be significant even when decoupled
from osmolality effects. How each variable affects the
growth phase and the production phase can be different depending upon the cell line, clone, and process. For example
in CHO cultures, increasing pCO2 only affected cell growth
rates minimally whereas, the impact of osmolality was quite
significant on cell growth rates. On the other hand, both factors had an impact on the production phase in terms of mAb
titers [160]. Some work has been published on the effect of
pCO2 and osmolality on NS0 cultures. These papers describe
different effects regarding the impact and conclude that there
are differences due to clonal variation and process design
[161-163].
As mentioned above, agitation is one of the variables
used to control DO and pCO2 levels. Historically, animal
cells were considered highly shear sensitive, but more recent
research confirms that animal cells are quite robust with respect to pure hydrodynamic forces [164-167]. Shear sensitivities are greatest at the air-bubble interface; therefore, the
presence of protective polymers in the culture medium is a
critical component. Typical additives are Pluronic F68 (most
common), albumin, and dextran [168], that help in protecting
the cells from high shear generated during bubble break up.
5.3.2. Nutrient Feeding
The simplest mammalian culture process is a batch process where all nutrients are added into the medium prior to
addition of the cells. The culture is maintained until cell
death and no further nutrient additions are made, on the assumption that sufficient nutrients are available in the basal
medium to achieve desired cell growth. While a batch process is simple and easy to implement, it is difficult to provide
sufficient nutrients in the basal medium without reaching
toxic levels of certain components or without generating
excessive levels of waste byproducts. A typical cell growth
profile in a batch cultivation shows a phase of exponential
growth which translates into a peak cell density that rapidly
decreases as cells quickly loose their viability and lyse. Researchers found that cell viability could be maintained over
an extended period of time, and even that a low level of cell
growth could be generated, if some nutrients were added at
different times during the cultivation. Initially, only simple
nutrients, such as glucose and glutamine were added as feeds
460 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
during the culture. Fed-batch cultures became more sophisticated as feed solutions were developed into more complex
mixtures [169]. One approach is to concentrate the basal
medium and feed the concentrated mixture at specified times
during the cultivation. Another approach is to analyze spent
medium to determine the consumption of medium components and design a feed solution based on this information
[144, 170-172]. While less empirical, the later approach can
be lengthy and may not yield a complete set of information
since cultivation media contain several dozens of ingredients, with some of them at difficult to measure concentrations. However, since some components may increase cell
mass while other components may increase specific productivity, a better understanding of the effect of individual components will allow different feed solutions to be designed for
different stages of the culture process and will support a
more economical cultivation process. Feed solutions can also
be designed to minimize undesirable byproduct generation
[173-178]. An obvious example is the substitution of galactose, a more slowly metabolized carbon source, for glucose,
which will support lower lactate production [173, 174, 177].
The majority of industrial mammalian culture processes use
fed-batch technology since with expertise, the feed solutions
and feeding regimens are reasonably simple to design and to
implement. Given the popularity of this feed approach, the
design of the production facility is simple and process development and validation are easy to perform. In addition, production phase process cycle times are reasonably short and
reduce process exposure to adverse events such as equipment
failure, and each fed-batch is easily associated with a product
lot.
At the extreme, a continuous feed can be provided to the
culture at a very low rate, where fresh medium is added to
the culture and spent medium (containing the product of interest) is removed from the culture. Both rates need to be
equal in order to maintain a constant volume. The advantage
of this continuous culture approach is that the cultivation
conditions are maintained constant over time and that waste
products are constantly removed. However, this approach
also results in the removal of perfectly viable cells from the
bioreactor. To improve on continuous cultivation methods,
the partial recycling of cells has been implemented and is
known as perfusion culture [179, 180]. Under these conditions, a pre-set cell concentration is maintained by returning
the desired amount of cells to the bioreactor. Perfusion processes are often maintained for many months, making it difficult to address regulatory issues on a per batch basis, and
requires purification of the product from very dilute harvest
streams. In addition, perfusion processes result in exposure
to adverse events over a long period of time, require large
amounts of expensive cultivation medium, and necessitate
the design, maintenance and operation of very complex and
difficult to scale up cell separation devices [181]. Also of
importance is the need to use a production clone that exhibits
excellent stability over several months of cultivation. For this
reason, this technology is seldom used. However, the benefit
of a perfusion process over a fed-batch process is that, for
the production of a similar mass of mAb, it generally requires smaller cultivation vessels and smaller factory footprints [181]. In addition, perfusion processes minimize product exposure to adverse culture conditions, and it is for this
Chartrain and Chu
reason that the one notable industrial exception is the production of Factor VIII where the product is highly unstable
[182]. However, since mAbs are notoriously stable once secreted in the medium, the implementation of the highly complex and sensitive perfusion technology may not be the most
efficient process. Fig. (4) pictures schematics and typical
growth, cell viability, and mAb production in batch, fedbatch and perfusion systems.
5.3.3. Cultivation Equipment
Stainless steel bioreactors remain the workhorse of the
industry. Stirred tank bioreactors used for production processes range from 1,000L to 25,000L in volume. These reactors are designed with baffles and a variety of impellers that
result in the desired mixing profile [183, 184]. Airlift bioreactors, ranging from 2,000L to 5,000L, are also used for production processes. These reactors are similar to a bubble
column where high air flow rates circulate upwards through
an inner tube and bubbles are released at the top of the tube.
The degassed liquid then flows downwards from the top of
the tube; thereby, creating circulation of the medium and
cells within the reactor [185, 186]. More recently, pilot-scale
disposable reactors have become available in volumes ranging from 50L to 2,000L. The use of disposable equipment
offers a wealth of advantages including the reduction of
preparation time, the elimination of cleaning and sterilization
steps, and a greater ease of use [187]. When factoring these
benefits, the cost savings in terms of time and capital are
likely to be significant. The manufacture of hundred of milligrams to gram quantities of recombinant proteins, produced
via animal cell cultivation, is often required in order to support animal and clinical evaluations. Generally, these support
activities rely on the cultivation of the animal cells in stirred
laboratory- and pilot-scale bioreactors. While highly reliable
and flexible, the preparation, operation, and cleaning of the
reactors are time consuming activities. However, the recent
commercialization of disposable and easy-to-use animal cell
cultivation devices, such as the Wave BioreactorTM (WBR),
Hyclone S.U.B., and Xcellerex XDR™, offer the prospect of
reducing the use of laboratory- and pilot-scale stirred bioreactors. In addition to their simplicity of use, the costs of disposable bioreactors and their ancillary accessories may be
lower when compared to that of a sterilizable-in-place bioreactor.
Briefly, the WBR consists of a sterile disposable plastic
bag that is half filled with cultivation medium with the head
space filled with the desired gas mixture. The bags are
placed on a rocking platform that delivers a wave-like motion to the liquid thereby delivering adequate mixing and gas
transfer to the culture while avoiding the formation of damaging gas bubbles [188]. WBRs offer the possibility for continuous gassing and are available in nominal volumes ranging from 2-L to 1,000-L (www.wavebiotech.com). Since
their market introduction, WBRs have been used for the cultivation of suspension [188, 189] and anchorage dependant
mammalian cells [190, 191], as well as insect cells [192].
Most recently, Hami et al. report the use of WBRs for the
clinical production of activated autologous T cells used in
the treatment of various forms of cancers [193]. WBRs have
also been fitted with a floating filter and successfully used as
perfusion reactors, supporting cell concentrations of up to
Development and Production of Commercial Therapeutic Monoclonal Antibodies
Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
461
Fig. (4). mAb production processes.
Overview of operations and typical cell viability and product production over process time. The average maximum cell densities are ~1-2 106
cells/mL for batch cultivation, 8-12 106 cells/mL for fed-batch cultivation, and 10-30 106 cells/mL for perfusion cultivation.
3x107 cells/ml, a six-fold increase over the cell concentrations routinely achieved in batch cultures [194].
The Hyclone S.U.B. and Xcellerex XDR™ are more
closely modeled after the stirred tank bioreactor. Both are
contained within a stainless steel outer shell that is not disposable and provides the temperature control. Disposable
sterile plastic containers are supplied from the vendor with
ports for typical pH, DO, and temperature probes. A sterile,
disposable impeller is built into each plastic container. The
agitation is bottom driven in the Xcellerex XDR and top
driven in the Hyclone S.U.B. The XDR™ is available from
200L to 2,000L (www.xcellerex.com), and the Hyclone
S.U.B. (www.hyclone.com) is available from 50L to 1,000L.
Minimum and maximum working volumes are similar to
stirred tank reactors where impeller location and mixing profiles determine these values. Both models have been used to
culture CHO and hybridoma cell lines, specifically for internal development projects at the respective companies.
Stainless steel stirred tank reactors still remain the industry standard, but disposable systems are gaining popularity
for specific uses, such as seed train expansion or material
generation. Since disposable systems do not require large
capital investment, smaller organizations may find them useful to create short term capacity. Fig. (5) compares the most
popular bioreactors currently in use for the cultivation of
mammalian cells.
6. SCALE DOWN CULTIVATION SYSTEMS
Traditionally, experiments were sequentially performed
in laboratory-scale bioreactors (i.e. 2L – 15L), scaled up to
pilot-scale bioreactors (i.e. 300L – 2,000L), and confirmed at
production-scale bioreactors (i.e. 5,000L – 25,000L). Today,
shortened project timelines, the vast possibility of therapeutic molecules, and the cost-conscious evolution of the indus-
try have forced process development groups to consider alternative scale down systems that have higher throughput,
lower costs, and better predictability of production-scale
bioreactors. Many candidate systems are available, but the
industry continues to evaluate the options. Bioprocessors'
SimCell system (www.bioprocessors.com) is one of the
more extensive and complex systems available. The SimCell
system consists of 700 μL reactors, 6 reactors per tray, where
up to 210 trays (1260 reactors) can be handled by the integrated robotics and data analysis system. This is a standalone system that is integrated with a control system, incubators, sampling systems, and some analytical systems. At the
other end of the complexity spectrum are well plate systems
and disposable tube systems (www.sartorius.com, Culti
Flask 50). These systems can be as simple as a shaker incubator and disposable well plates or 50 mL tubes that are
manually inoculated, sampled, fed, and maintained with little
to no on-line monitoring. Feeding, sampling, and harvesting
can be automated by setting up a liquid handling system with
an automation control system that can be programmed by the
individual users. Applikon micro reactors (www.applikonbio.com) are similar to well plate systems but with additional
sensors and control systems included. The micro reactors
look similar to 24-well, deep-well plates, but each plate contains sensors, which the automation system uses to control
pH, temperature, and DO levels; although, currently, the
system only has one-sided pH control for mammalian cultures (CO2-based control). The micro reactors were originally designed for microbial cultures using CO2 and ammonia gas for pH control, but these systems do show potential
for mammalian cultures [195]. A common theme that scale
down systems share is the use of alternative pH and DO
probes – particularly fluorescent (optical) probes – instead of
traditional electrochemical probes [196]. As a direct result of
fluorescent technology, smaller, less invasive, disposable
462 Current Pharmaceutical Biotechnology, 2008, Vol. 9, No. 6
Chartrain and Chu
Fig. (5). Typical vessels used for the cultivation of mammalian cells.
Fig. (6). High level overview of the different activities leading to the implementation of a mAb production process at industrial scale. The
timelines indicated are averages of what has been reported in Industry and can vary with the approach taken by each company and on the
individuality of each mAb expressed.
probes that require little to no calibration (probes are delivered pre-calibrated) are now available in the form of optical
probes. Optical probes are patches that are <1 mm in thick-
ness and can be adhered to the inside of a clear reactor surface, and the optical instrumentation is external to the reactor, which results in a non-invasive system. The fluorophores
Development and Production of Commercial Therapeutic Monoclonal Antibodies
are immobilized on the patches, and in the case of DO
probes, the emission of the fluorescent dye is quenched at a
rate directly related to oxygen concentration. pH probes, on
the other hand, use a fluorescent dye with an absorbance
spectrum that changes as a function of pH.
Many scale down systems, ranging from highly complex
to simple, are available for evaluation. As the field matures,
we will see which systems are best for prediction of production-scale performance.
[12]
[13]
[14]
[15]
[16]
CONCLUSION
In this review of therapeutic mAb production by cultivation of mammalian cells, we have tried to provide the readers
with an encompassing overview of the field. From the fundamentals of mAb biochemistry that drives many decision
made during development to the current state of large scale
cultivation of the engineered cells. References to the ongoing
introduction of novel technologies, from advances in cell line
engineering to the introduction of disposable bioreactors
were provided to give the readers a sense of the direction the
field is moving to.
In addition, we hope that we communicated that the discovery, development, and commercial production of therapeutic mAbs is a lengthy, scientifically and technically complex, and resource consuming process. Fig. (6) presents an
overview of these steps and their associated time lines.
At this time, the cost of therapeutic mAbs reflects these
economic burdens encountered during development and production, and has been challenged in many public forums
[197]. Having taken root in the late 1980s, this industry is
still very young and hopes are that continued development of
more efficient technologies such as these described in this
review will help in controlling cost of goods and will make
mAbs available to an ever increasing number of patients.
[17]
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[21]
[22]
[23]
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Received: April 29, 2008
Revised: June 03, 2008
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