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Файл:Ординатура / Хирургия / Библиотека им академика М.И. Перельмана / Книга_101_библиотеки_им_акад_М_И_Перельмана
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Downstream Processes
11
Involved in Protein
Production
11.1 OVERVIEW
The crude protein originates in the upstream process, where conditions inuence protein type, but
the downstream process denes the nal product. The therapeutic product, such as secretion in
the media (mammalian cell culture) or microbial cell biomass or components, is obtained postfermentation. Downstream processing involves separation, recovery, and purication of the product
from the host system. Typically, the transition from upstream to downstream occurs during the
recovery step, i.e., harvest (claried supernatant or cell pellet). Regardless of the expression system,
cell removal is achieved through centrifugation or ltration. The chapter discusses the harvest of
culture broth, essential for recovery and purication of biological products, including insights into
future trends.
The chapter primarily focuses on the two most utilized production systems, Escherichia coli and
CHO, outlining purication and product concentration strategies for these systems before nal formulation and ll- nish of drug substance (DS)/ drug product (DP; Figure 11.1).
The initial step in selecting downstream processing methods involves understanding the protein’s
features, stability prole, and factors potentially affecting its structure. Downstream processing
schemes typically comprise capture, intermediate purication, and polishing stages (Figure 11.2),
each serving specic purposes. The capture step, the rst chromatographic unit operation, aims
to isolate and stabilize the target protein while removing key host cell components. The secondphase of purication eliminates most contaminants such as nucleic acids and host cell proteins; its
inclusion depends on succeeding chromatography processes and the purity prole of the target protein. The nal stage, polishing, aims for maximum product purity while eliminating product- related
impurities such as oxidized or deamidated species. The selection or combination of techniques
considers the target protein, product quality, and specic requirements.
The overarching workow and objective of a downstream process remain constant: ensuring the
product meets specied quality attributes and purity proles, establishing the product prole, and
determining critical product quality attributes during the initial phases of process development and
design. Purication and efcient recovery are pivotal factors contributing to the success of both the
upstream and overall bioproduction processes. The selected purication process needs to be robust,
scalable, and capable of effectively removing or reducing both product- related and process- related
contaminants to achieve high purity and ensure safety. Maintaining stringent quality standards for
recombinant proteins is paramount due to concerns about potential immunogenic effects or other
hazards arising from the product’s deviation from human- like characteristics or potential contamination by the host cell system (e.g., mammalian cells).
Selection of appropriate chromatography separation methods relies on differences in the properties of the target protein and other substances present in the sample. Chromatography, a fundamental
technique in protein purication, continually evolves as a high- resolution method. Both traditional
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DOI: 10.1201/9781003392026-11
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FIGURE 11.1 Overview of downstream unit operations for recombinant protein purication
FIGURE 11.2 A three- stage purication strategy
and advanced techniques leverage the physical and chemical properties of the protein, its interactive
nature, and the characteristics of product variants. Examples of protein characteristics used in
various chromatography methods are illustrated in Table 11.1 and Figure 11.3.
When choosing a resin, consider qualities such as dynamic binding capacity (DBC), yield,
quality, HCP removal, and purity.
Maximizing productivity and cost efciency involves minimizing the number of processes
without compromising overall yield. Conversely, downstream processing steps are critical for
product protection and therefore require close monitoring. The primary aim of purication is to
reduce or eliminate impurities and contaminants to levels that are safe for patient use. Impurities
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TABLE 11.1
Downstream Process
Protein Property and Applicable Chromatography Methods
Target Protein Applicable Chromatography Purication Method
Specic ligand recognition Afnity chromatography (AC)
Metal ion binding Immobilized metal- afnity chromatography (IMAC)
Charge Ion exchange chromatography (IEX)
Hydrophobicity Hydrophobic interaction chromatography (HIC)
Size Gel ltration (size exclusion chromatography)
Isoelectric point Chromatofocusing
FIGURE 11.3 Schematic presentation of the various chromatography methods
inuencing downstream purication strategy fall into two categories: process- related and product-
related impurities in biological manufacturing.
Product- related impurities in the drug substance are molecular variations that occur during
manufacturing or storage, differing from the intended products in terms of function, protection,
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or efcacy. Structural differences in proteins can arise from the cellular mechanisms of the host
organism in protein synthesis or from specic processing activities. These deviations may or
may not be acceptable from a patient safety perspective. Product variants are substances with
attributes (activity, safety, efcacy) comparable to the target product. Detailed characterization of
the product and its variants is necessary to differentiate between contaminants and product- related
chemicals.
Process- related impurities are contaminants associated with the manufacturing process, including
proteins from the host cell, DNA, and raw materials (e.g., cell culture media, antibiotics, chromatographic media used for purication, and specic buffer components). Upstream process impurities
may originate from the cell substrate or the cell culture, inuenced respectively by the expression system and upstream process parameters that dictate whether the product is secreted into the
medium or remains intracellular. Downstream process conditions and source materials also affect
impurities, including chromatographic media, buffer components, and leachable components.
The primary objective of the downstream process is to diminish contaminants through a series of
purication procedures to ensure the nal product meets purity criteria.
Monitoring downstream unit operations involves screening the product at each stage for its purity
prole, stability, and biological activity. Emphasis on characterization, stability, and overall product
prole intensies as the target molecule progresses through developmental stages, from toxicology
to clinical and safety and efcacy testing. Testing commences only after nalizing the upstream
and downstream processes, underscoring their pivotal role in the biopharmaceutical development
timeline.
11.2 E. COLI SYSTEM: RECOVERY AND PURIFICATION
Recombinant protein production in E. coli occurs in three spaces: intracellular, periplasmic, or extracellular. However, the primary site for production is intracellular, resulting in insoluble aggregates
known as inclusion bodies (IBs).
Inclusion bodies, dense protein clumps found in both the cytoplasm and periplasm, possess a
higher density than cellular debris, aiding their separation during the process. Proteins within IBs
remain generally stable and serve as a suitable stage for halting manufacturing operations. These
bodies often contain more than 90%– 95% of the protein, lessening the burden of downstream purication. Yet, the protein remains inactive and insoluble, necessitating conversion into its active, native
conformation. Once achieved, the functional protein undergoes purication and concentration for
nal product formulation.
Steps preceding purication involve converting the insoluble protein form to its soluble,
active form
• The cells must be broken to release the inclusion body.
• The inclusion body is separated from the cell detritus.
• Co- precipitated pollutants are removed by washing the inclusion body.
• The inclusion body is solubilized to denature it in preparation for refolding.
• The process of restoring a protein to its active state is known as refolding.
Recovering the inclusion body and renaturing the protein into its native state pose additional
challenges in the processing stage.
Figure 11.4 depicts a typical downstream process for a recombinant protein generated as inclusion bodies in E. coli.
For the expression of soluble proteins in the E. coli cytoplasm, inclusion bodies do not form, simplifying the process to recover the desired protein. Certain cytoplasmic proteins can spontaneously
fold under optimal conditions.
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FIGURE 11.4 Flow chart for downstream processing of recombinant protein expressed in E. coli
Modifying the expression mechanism and appropriate vector design at the cellular level redirect
intracellular protein expression from inclusion bodies to soluble cytoplasmic proteins. While soluble protein expression is an attractive option, its suitability varies among protein types. The success
of this strategy relies on expressing the protein abundantly, at least exceeding 5% of the total protein,
enabling maximized recoveries without additional steps (IB recovery, solubilization, and refolding).
11.2.1 cell disRuPtion
Cell disruption to extract inclusion bodies (recombinant protein) occurs after cell harvest during intracellular protein synthesis in E. coli. E. coli’s double cell wall consists of an outer lipopolysaccharide
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(LPS) membrane, an aqueous periplasmic gap, and an inner thin peptidoglycan cell wall. The
breaking of these cellular membranes for accessing the intracellular area is known as cell lysis
(involving the outer LPS- rich membrane and the cytoplasmic inner membrane). Cell disruption
methods include chemical, enzymatic, mechanical, or a combination thereof.
The appropriate method is determined based on the protein of interest, the technique’s impact on
product quality, downstream processing, and overall yield. Additionally, considering the product’s
characteristics (solubility, cellular location, and physical properties), batch size, operational scale,
processing time, efciency, and the protein’s stability is crucial for evaluating and determining the
optimal method.
11.2.1.1 Physical Methods
Various methods, such as mechanical disruption (e.g., grinding, high- pressure homogenization, and
ultrasound), and non- mechanical disruption (e.g., freeze/ thaw, heat treatment, and osmotic shock),
are utilized to physically lyse cells.
11.2.1.1.1 Mechanical Methods
Mechanical disruption methods are preferred in industries due to their high disruption efciency and
scalability. These methods involve handheld grinders or motorized devices with rotating blades that
break down cells. Common devices include the Waring blender, Polytron, hand- held grinders such as
Dounce and Potter- Elvehjem homogenizers, and high- pressure homogenizers like the French press.
The Waring blender resembles a typical household blender and is primarily used for grinding.
The polytron operates by drawing tissue into a long shaft with rotating blades. Both these devices
excel at grinding soft, solid tissues. The Dounce homogenizer functions by pushing the sample
between the tube’s sides and the pestle, generating shearing forces. The Dounce homogenizer is
an effective for lysing tissue culture cells, ne tissue pieces, or applications requiring only mild
lysis. Conversely, the Potter- Elvehjem homogenizers grind tissues and also lyse cells. The sample
is placed in the sample tube, and the pestle is rotated at a certain speed. This repeated movement of
the pestle on the tube causes a shearing force that breaks the cells.
However, the most prevalent industrial device is the high- pressure homogenizer. Samples pass
through a narrow space while experiencing high pressure due to piston action (e.g., 20,000 psi),
causing cells to expand and rupture as they transition from high to low pressure zones. Typically,
samples undergo multiple passes through this homogenizer for enhanced lysis efciency. To reduce
protein denaturation, it’s advisable to cool the sample (2– 8ºC) during homogenization. Highpressure homogenizers offer scalability and accommodate various sample volumes (Figure 11.5).
A bead mill homogenizer is a commonly used mechanical method for disrupting E. coli. This
method involves beads that are vigorously agitated at high speeds to break and lyse the cells. The
cell suspension passes through a grinding chamber (which contains a rotating shaft) lled with
beads (approximately 80%). When exposed to high speeds, the sheer force and impact from the
beads cause the cells to break. The device does not use any external probes and is relatively selfcontained, minimizing contamination risks.
Sonication is another mechanical disruption method used for cell lysis. This technique employs
an acoustic transducer to generate pulsed high- frequency sound waves, creating microscopic bubbles
that radiate through, leading to agitation and eventual cell lysis. It is useful for homogenizing small
sample volumes and effectively targets bacteria and spores. However, excessive heat is typically
generated from the ultrasonic treatment, so samples immersed in an ice bath are usually subjected
to short bursts.
Despite being commonly used, mechanical disruption methods have challenges. These include
heat generation, potential denaturation of proteins, limited selectivity, shear stress that might damage
sensitive components (such as specic proteins), and the disintegration of cell debris, making the
recovery process more prone to contamination.
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FIGURE 11.5 Schematic of the working principle of a high- pressure homogenizer
11.2.1.1.2 Nonmechanical Physical Methods
Freeze/ Thaw The freeze/ thaw method involves freezing a cell suspension (using dry ice, ethanol,
or a freezer) and then thawing the cells at ambient temperature or at 37ºC. Ice crystals form during
freezing, causing cells to expand and lyse during thawing. As a result, multiple freeze/ thaw cycles
are necessary for effective lysis.
Heat Shock Thermolysis uses heat to denature membrane proteins and lyse cells at temperatures
approximately 50ºC for outer membranes or 90ºC for cytoplasmic components.
Osmotic Shock Cell lysis using this method involves a signicant alteration in the osmolality of the
cells. Initially, cells are immersed in a high osmotic pressure solution (e.g., 1 M sucrose or a highsalt solution; 1M). This is swiftly transitioned to a low concentration solution, inducing osmotic
shock within the cells. Consequently, water molecules move from the low- salt concentration to the
high salt concentration (into the cells), causing increased pressure within the cells and resulting
in an explosion. Conversely, exposing cells rst to a low concentration solution and then to a high
concentration solution leads to the outow of water from the cells, causing rupture.
11.2.1.2 Chemical Methods
In addition to mechanical and non- mechanical approaches, chemical disruption methods involving
enzymes or chemical additives such as detergents are also utilized to lyse cells. These agents work
by enzymatically destroying the cell wall or membrane, increasing osmotic pressure, or precipitating cell wall proteins.
Detergents function by forming micelles, binding to the lipid membrane protein, and causing the
rupture of the cell membrane, enabling internal proteins and inclusion bodies to escape. Detergents
vary based on physical properties and action type:
• Anionic: Sodium dodecyl sulfate (SDS) reorganizes the cell membrane by disrupting proteinprotein interactions.
• Cationic: Ethyl trimethyl ammonium bromide may disrupt LPS and phospholipids in cell
membranes.
• Triton X100 is a non- ionic detergent that solubilizes membrane proteins.
Solvent addition, chaotropes, and metal chelators are other chemical additives used for cell
disruption. Solvent addition works by extracting the lipid components of the cell wall, increasing
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cell membrane permeability, and releasing intracellular proteins. Commonly used solvents include
toluene, dimethyl sulfoxide, and certain alcohols. Chaotropes such as urea can solubilize membrane proteins. EDTA, a chelator, binds to metal ions Mg
2+
and Ca
2+
on the cells’ outer membrane,
increasing permeability and causing the release of intracellular components.
The use of digestive enzymes that disintegrate the cell wall is another chemical technique for
lysing cells. Enzyme selection depends on the variability of cell walls and membranes among
different cell types and strains. For instance, lysozyme is commonly used to break down the cell
walls of gram- positive bacteria.
The chemical method is easy to implement and scalable compared to mechanical methods but
requires substantial capital expenditure. However, limitations include prolonged reaction times for
efcient lysis, downstream operations’ burden to ensure effective chemical removal, and potential
hazards posed to operators by these chemicals.
Typically, a combination of disruption methods is employed in biomanufacturing to overcome
efciency limitations. For example, using chemical detergents (Triton X- 100) as a pretreatment
followed by mechanical disruption (high- pressure homogenization) enhances lysis efciency and
reduces passes through the homogenizer. Similarly, chelating agents and detergents are included in
buffers to suspend the cell pellet before liquid homogenization, improving efciency compared to
using a single method.
In the preparation of soluble proteins expressed in E. coli, cells undergo lysis, and the supernatant
is cleared to eliminate cell debris, lipids, ribosomes, and other particulate materials. The cleared protein is then introduced to the appropriate chromatographic column for capture.
11.3 INCLUSION BODY RECOVERY AND SEPARATION
After cell rupture and the release of their contents, including inclusion bodies and other cellular
components, recovery and separation processes are carried out. These processes are essential for
separating insoluble inclusion bodies from soluble components and insoluble cell debris.
Various separation procedures based on inclusion bodies’ density and solubility variations have
been applied to increase yield and purity. Clarifying, purifying, and concentrating inclusion bodies
before solubilization and renaturation require inclusion body recovery and separation. Methods such
as centrifugation, ltration, or alternatives such as precipitation and two- phase aqueous extraction
may achieve relatively pure inclusion bodies.
Centrifugation is a commonly employed technique for inclusion body recovery. Benchtop/ oor
and continuous centrifuges may be used depending on the operation’s scale and liquid volumes.
Adjusting speed and gravitational force (usually between 5,000– 20,000 g) allows collection of
inclusion bodies as pellets, while the less dense cell debris is removed in the supernatant, reducing
the processing volume.
Microltration or ultraltration is an alternative approach for the recovery of insoluble inclusion bodies. Inclusion bodies are between 50 and 1500 nm in size, and choosing an appropriate
large pore size lter (microltration: 0.05 µm to 5 µm, ultraltration: 0.001 µm to 0.1 µm)
achieves a high yield of inclusion bodies. Additionally, ltration aids in enhanced purication by
removing soluble impurities, DNA, and toxins released during cell disruption through the large
pore size.
Microltration is used to remove cells and cell detritus from the cell lysate, targeting a size range
of 0.2 μm to 10 μm. Ultraltration membranes, with pore sizes spanning from 0.001 μm to 0.1 μm,
effectively eliminate larger molecules like proteins, polysaccharides, and IBs. To address impurities
potentially co- precipitating with inclusion bodies, a dialtration step employing an inclusion body
wash buffer can aid in their removal. However, its practicality for large- scale operations is limited
due to the substantial wash solution consumption required for efcient processing. Moreover, it’s
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Downstream Process
advisable to maintain a low transmembrane pressure (TMP) during cell lysate processing, with consideration for factors such as processing time, impact on product quality, membrane fouling, and
performance. Essential parameters for ltration operations include membrane pore size, membrane
type (hollow ber, at sheet), permeate ux/ fouling, TMP, and cleaning.
In addition to standard centrifugation or membrane ltration, other methods like expanded bed
adsorption, aqueous two- phase liquid extraction, solvent extraction, precipitation, and size exclusion chromatography offer alternative approaches for isolating and purifying inclusion bodies from
cell lysate, integrating cell disruption and solubilization.
The selection of the most suitable method— centrifugation separation, membrane ltration, or a
combination of cell disruption and solubilization approaches— depends on various factors, such as
the protein of interest, operational scale, facility design, buffer consumption, overall production process, and desired purity levels. It is equally crucial to assess the challenges and limitations of each
method to ensure reliable, robust, and economically feasible recovery and separation of inclusion
bodies.
Depending on the chosen approach, integrating wash solutions might be necessary due to the
highly insoluble nature of inclusion bodies, particularly to eliminate cellular impurities that coprecipitate with them. Despite the generally high purity of most inclusion bodies, optimizing wash
solutions signicantly enhances overall purity and streamlines operations. Enhancing purity before
solubilization and refolding of inclusion bodies can lead to increased overall process yield and
potentially reduce subsequent processing volumes.
For instance, recovered inclusion bodies may undergo washing with solutions containing specic components (e.g., low levels of chaotropes, detergents, salts, and pH adjustments) to effectively remove cell debris and other co- precipitated impurities. Striking a delicate balance between
purity and recovery is vital when selecting chaotropes or agents that could potentially impact inclusion body solubilization and overall yield (e.g., using high concentrations of urea or guanidinehydrochloride). Typically, centrifugation is employed to recover inclusion bodies post- washing.
Finally, puried water serves as the last- step wash, removing chemical agents and debris before
yielding a highly pure inclusion body pellet.
At this stage, the concentrated and pure inclusion bodies (IB) can be frozen for long- term storage
(typically at −20ºC). This allows for exibility in production schedules, enabling the pooling of
IB pellets from multiple upstream batches for further processing and maximizing downstream
operations.
11.3.1 inclusion body solubilization
Once the IB pellet is recovered, a crucial step is to re- solubilize it in preparation for subsequent
renaturation. The aim of the solubilization process is to denature the protein and decrease the amount
of aggregated protein in the inclusion body. This denaturation of the insoluble protein aggregate
aims to transform it into a soluble linear polypeptide, achieved through a combination of chemical
agents and specic operating conditions.
Traditionally, substantial amounts of a denaturant have been used for solubilizing inclusion
bodies. Common substances include 6 M guanidine- hydrochloride (GuHCl), 8 M urea, detergents,
alkaline pH (>9), organic solvents, or N- lauryl sarcosine. Gu- HCl, being a potent chaotrope, is
notably more effective than urea at denaturing excessively aggregated inclusion bodies. However,
urea solution containing isocyanate can lead to carbamylation of free amino groups of polypeptides,
particularly when exposed to prolonged incubation under alkaline pH, impacting product quality.
A challenge with high concentrations of chaotropes is the possibility of precipitation, affecting
overall process yield and subsequent purication steps.
Before incorporating the chosen denaturant into the solubilization technique, it is crucial to test
it with the target protein.
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TABLE 11.2
237
Example of Solubilization DoE
Process Parameters Start Condition Optimization
Buffer 50 mM Tris- HCl, pH 8.0 pH (7.0, 7.5), buffer composition (e.g., EDTA)
Denaturant 8 M urea or
6 M Gu- HCl
Inclusion body concentration
(mg/ mL, wet weight)
Temperature (°C) Ambient 4– 30
Time 60 min 15 min to 2+ hours
20 Testing higher and lower concentrations (e.g., 10 and 40 mg/
Concentration of the denaturant
mL)
Factors such as the presence and concentration of a reducing agent, as well as time, temperature,
ionic strength, and the denaturant- to- protein ratio, signicantly inuence the success of solubilization for each denaturant. Table 11.2 provides the initial experimental conditions for solubilizing
inclusion bodies, often followed by purication compatible with the solubilized proteins.
Numerous solubilization techniques (e.g., REFOLD database: http:// pford.info/ refol ddb/ ) have
been published, including the use of SDS (10%), N- lauryl sarcosine, or other detergents, and extreme
pH levels as alternatives to the commonly used solubilization agents. For proteins with disulde
linkages, reducing agents like dithiothreitol (DTT), beta- mercaptoethanol, or Tris 2- carboxyethyl
phosphine (TCEP) are commonly employed to decrease disulde bonds and stabilize free cysteines.
Milder solubilization solutions with lower chaotrope concentrations have been used for some
proteins. Employing mild solubilization conditions has shown to retain native- like secondary
structures, potentially improving protein renaturation by denaturing the protein to an intermediate
state instead of completely linear polypeptides. This approach might enhance refolding yields, but
its applicability heavily depends on the protein and the produced inclusion bodies.
Solubilization signicantly impacts the subsequent refolding conditions; hence, the solubilization
buffer and operating conditions must align with the renaturation process. For proteins containing
cysteine, isolated inclusion bodies often possess non- native intramolecular and intermolecular disulde bonds. Incorporating reducing reagents with chaotropes can aid in reducing these bonds. The
non- native disulde bonds can likely reduce the solubility of the IBs in the absence of dithiothreitol,
cysteine, or beta- mercaptoethanol. The inclusion of thiol agents is unnecessary for proteins that may
already be in a reduced state. Arginine is commonly used in solubilization and/ or refolding buffers
to prevent protein aggregation, while EDTA may be employed to prevent oxidation of free cysteine
groups exposed during denaturation and reduction.
Apart from the denaturant type and additive components, parameters such as reagent concentration, solubilization buffer- to- IB pellet ratio, reaction time, pH, and temperature must all be evaluated
for optimal solubilization conditions.
11.3.2 inclusion body RenatuRation
Protein refolding begins with the solubilization of inclusion bodies (IBs) to acquire properly folded
proteins. Gradual elimination of excess denaturants and reducing agents occurs, and reduced proteins
are then transported to an oxidizing environment, initiating the formation of native disulde bonds.
Renaturation of the solubilized IBs is achieved through dilution, dialysis, or on- column processing.
Typically, refolding is conducted at very low protein concentrations, usually approximately 10– 100
g/ mL, to prevent aggregation and gradually convert denatured proteins to their native, properly
folded structure.
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