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particles. Choosing a lower ow rate can provide sufcient time for smaller particles to settle
before leaving the rotor or bowl, and optimizing ow rate can enhance process efciency.
• Cell density, viability, and cell type are other variables to consider when choosing centrifugation as a separation choice. The solid to liquid ratio will determine how soon the pellets (solid’s
settling) accumulate and ll the bowl leading to a possibility of packing up into the claried
stream. As such, the pellet may need to be intermittently or continuously removed. Turbidity
measurements of the supernatant/ claried liquid can be useful in such scenarios, which will
increase as the bowl reaches its holding capacity. A balance is critical to minimize loss.
Assessing these variables on scale- down models can be challenging due to the limited availability of suitable representative models for continuous ow centrifuges. Benchtop or conventional
centrifuges cannot replicate the dynamics of a continuous centrifuge.
10.6.2 dePth filtRation
Unlike surface or membrane ltration, depth ltration employs the depth of the ltering medium and
its absorptive properties (porous substrate) to retain particles from the feed stream. The structure of
depth lters consists of multiple layers or even a single thick layer of culture media. Larger particles
are captured at the surface, while smaller or ner particles are entrapped within the various layers or
structures. Depth lters are not identied to have a dened pore structure like other lters do. In a
membrane lter, particles larger than the pore size are trapped on the membrane’s surface. Thus, the
depth lter leverages the depth of the membrane media and surface- based sieving for applications
such as cell culture harvest and intermediates (e.g., precipitated solutions, refolded protein processing before chromatography steps; Figure 10.8).
FIGURE 10.8 Comparison of membrane and depth lter functionality
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Depth lters are constructed using cellulose or polypropylene bers combined with lter aids
like diatomaceous earth (DE) or perlite. These bers are bound together with a polymeric resin
using the wet- laid process, forming the matrix of the lters. The ber matrix provides rigidity and
structure, while lter aids increase surface area to enhance retention properties. Additionally, the
resin imparts wet strength and a positive charge to the lter surface. Some lters may incorporate an
additional lter membrane layer with different pore sizes (e.g., 0.2 μm). When selecting a suitable
depth ltration system for primary or secondary clarication of the feed, consider variables such as
lter selection, format, and surface area. The choice of depth ltration for cell culture applications
depends on culture conditions and the physical properties of the product, particularly if it involves
proteins present in the feed for mammalian cultures.
Depth ltration offers several advantages, including ease of implementation, single- use capability, faster turnaround times, reduced or no cleaning requirements, scalability, and low investment. In single- use systems, lter units can be stacked together to increase surface area, allowing
for easy scaling to handle large volumes (greater than 5,000 L of harvest). These lter units, or
capsules, consist of lter culture media, along with inlet and outlet single- use manifolds equipped
with vents positioned between lters. These capsules are arranged into stainless steel holders,
although the stainless steel parts do not come into contact with the product. Commercially available single- use systems, such as 3M Encapsulated Zeta Plus and Millistak+ Pod lter systems by
Millipore, are widely used in various industries. Depth lters are well- suited for a range of clarication applications, including cell culture, yeast, E. coli lysates (after centrifugation), refolded proteins
(after E. coli IB solubilization), vaccines, and plasma proteins.
The setup for depth ltration is straightforward (Figure 10.9). Before starting the harvest process, it’s essential to ush the depth lters with water or a recommended buffer to remove any
loose particulates and extractives that may have been trapped during the manufacturing process.
Subsequently, the unclaried culture is introduced into the lter setup through the inlet. A pump is
then used to drive the feed from the outside, passing through the lter culture media and traveling
through its depth to achieve clarication. After completing the harvest process, the lter culture
media is once again ushed with water buffer to recover the unit’s volume and minimize product loss.
The void volume of depth lters varies depending on pore structures, with more open pore
structures having a high void volume (around 90 percent) and tighter pore structures having lower
void volumes (around 30– 40 percent). To ensure successful clarity, it is crucial to select the appropriate lter grade based on the particle size distribution. In many cases, multiple lter grades are
arranged in series to improve overall clarication performance. Following clarication, the harvested
material is ltered through a 0.2 μm lter. Single- use bags are advantageous in this application for
easy storage and maintaining sterility until further processing. Properly sizing the depth ltration
process is essential for tight process control, preventing batch losses, or processing delays. Depth
ltration also offers benets in reducing shear stress, limiting cell damage, and removing impurities
such as HCP, DNA, and endotoxins, compared to centrifugation.
10.6.3 ultRafiltRation and MicRofiltRation tangential floW filtRation
Membrane ltration techniques, such as ultraltration and microltration, can be effectively
used in the recovery and separation process post- fermentation. Hollow ber lters have enabled
FIGURE 10.9 Schematic of a depth ltration setup (primary or secondary clarication)
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the application of ultraltration and microltration TFF (tangential ow ltration) in large- scale
operations, including clarication and cell retention in perfusion processes. The choice between
microltration and ultraltration depends on membrane pore size, with microltration typically
having pores around 0.1 μm and ultraltration membranes having even smaller pores categorized by
molecular weight cutoff. While microltration and ultraltration are commonly used in downstream
processing, recent advances have positioned these methods as suitable alternatives to traditional
centrifugation. TFF is successful in cell harvest and cell lysate processing, relying on size exclusion
for ltering separation and utilizing crossow to remove cells and detritus from the membrane’s surface. The retentate pool containing cells is concentrated while the cell culture is treated. A frequently
encountered issue with this system is lter fouling, which drops the performance and recovery.
This can be overcome by incorporating dialtration or cell washing, which is benecial for buffer
exchange in chromatography applications and cell lysis.
The selection of the appropriate membrane pore size is crucial for process efciency. Hence,
identifying the membrane with the appropriate pore size is key to developing an effective clarication or cell harvesting process. Smaller pore size membranes provide higher permeate ux under
steady- state conditions, so choosing a larger pore size ultraltration membrane for E. coli, even with
relatively small pores compared to the cells, may be advantageous. For applications where proteins
are secreted in the medium, open pore size microltration membranes in the range of 0.2 to 0.65
μm are preferred, especially for larger recombinant proteins like monoclonal antibodies. Selecting a
membrane pore size at least ten times larger than the target material ensures proper passage through
the membrane.
In the case of hollow ber lters, the cartridge’s length and the inner diameter of the bers
inuence the process. Shorter path lengths are suitable for processing cultures. Other variables
affecting the clarication process include permeate ow rate, recirculation ow rate, and timing of
dialtration (cell washing) to facilitate particle passage (cells or proteins).
10.6.3.1 Choosing the Appropriate Method
The primary goals for clarication include
• consistent removal or separation of cells,
• removal of insoluble debris,
• reproducibility and scalability
In addition to these goals, increasing throughput to reduce processing time, minimize product
degradation, product loss, and downstream operation burden are essential considerations when
selecting a clarication method.
The choice of clarication method should be based on the cell type. E. coli is primarily used for
producing simple proteins and antibody fragments that do not require glycosylation. Most bacterial
expression systems involve a centrifugation step to recover the pellet from which the protein is puried. Subsequently, cell lysis is performed to release the recombinant protein. During the cell lysis
and inclusion body (IB) recovery process steps, alternatives to centrifugation such as microltration
and ultraltration, can be considered for processing cell lysates and recovering IBs. For refolded
proteins, particularly on an industrial scale, depth ltration is often the preferred choice of clarication. Microltration membranes can concentrate cells and disrupt them using a homogenizer
to separate the inclusion bodies. Cell debris and IBs can be separated using either microltration
membranes (0.2 µm) or ultraltration membranes.
Harvesting methods for E. coli secreting soluble proteins are similar to those used in mammalian
production systems, such as centrifugation, depth ltration, and sterile ltration. While process
modications may be necessary to accommodate the more intensive fermentation characteristics
of E. coli cultures, like high cell density and short cell culture time, similar process equipment and
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FIGURE 10.10 Alternate ltration system for processing E. coli products
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optimization strategies should be expected. In processes where cells are engineered (e.g., using
secretion tags and other changes to the E. coli construct) to release the product from the periplasm
or into the extracellular space, it’s crucial to avoid shear stress on the cells. Precipitation and occulation (e.g., with polyionic polymers like PEI and dextran) in conjunction with centrifugation can
assist in rapid processing, stabilizing the product, and efciently removing cell debris, colloidal
proteins, and nucleic acids.
CHO cells are another common expression system. In most mammalian systems, proteins
are secreted from these cells into the culture medium. Therefore, the clarifying process involves
removing cells and soluble debris. Centrifugation, depth ltration, or tangential ow microltration
are commonly used for primary clarication. To eliminate smaller particles and protect the chromatography column, a nal ltration step is included. For this purpose, a 0.2 µm rated sterilizing
grade lter is used. To preserve the more sensitive sterilizing grade lter layer, a dual- layer sterile
lter, such as a 0.45 µm over a 0.2 µm lter, is commercially employed. Recent techniques, such
as intensication and cell engineering, have led to higher cell densities, necessitating improved
impurity removal to facilitate downstream operations. The clarication process is critical to minimize particle content (cells, debris, impurities like DNA, endotoxin, HCP) in the load for the subsequent chromatography step (e.g., Protein A) and reduce fouling of the stationary phase.
The solid content per unit volume, often referred to as cell density and viability, plays a crucial
role in determining the choice of clarication method, performance efciency (yield), and overall
effectiveness. High cell density can lead to increased levels of DNA, HCP, and other cell debris.
Centrifugation is effective for handling high cell concentrations and processing large volumes,
rendering depth ltration or microltration less effective at such high cell densities. However, centrifugation can have drawbacks due to the shear stress it imparts on cells, increasing the risk of
cell lysis and impurities. To address this concern, occulation or precipitation techniques may be
employed. Figure 10.10 illustrates the available options for establishing a robust harvest system
tailored to the specic cell line and target protein.
The modernization of existing harvesting methods to accommodate high cell density, increased
biomass, and improved impurity removal can signicantly enhance the purication process.
Therefore, it may be benecial to explore additional techniques and strategies, if necessary, to
achieve the highest level of clarication efciency.
10.7 ANALYTICAL TOOLS
Qualitative and quantitative analytical tools play a pivotal role in validating the process across
various scales. Analytical methods must be precise, accurate, and robust. The data obtained from
these analyses inform decision- making throughout the entire process, from early- stage cell line
development to the nal product. In- process testing is a crucial component in enabling Process
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Analytical Technologies (PAT), which involves understanding process parameters, conditions, and
materials to develop an effective control strategy. According to the ICH Q10 guideline, a control
strategy is dened as “a planned collection of controls derived from current product and process
understanding that guarantees process efciency and product quality.” Here, we summarize some of
the critical upstream analytical tools frequently used.
10.7.1 cell cultuRe/ feRMentation
Growth monitoring is the primary analysis in the culture process, determining whether the existing
conditions favor cell growth. Various ofine methods are available for monitoring bacterial cells, with
spectrophotometry being the most widely used due to its simplicity, speed, and reliability. Microscope
counting is an alternative, albeit cumbersome and less reliable method. For mammalian cell cultures,
the trypan blue method (using a microscope and hemocytometer) is common. Several automated
cell- counter instruments, such as the Vicell Cell Viability Analyzer and Cedex Cell Analyzer, offer
pre- programmed and customizable solutions, estimate cell sizes, and handle various animal cells.
While these methods remain popular for cell density measurements, recent years have seen the
emergence of online cell density measurements, similar to the monitoring of pH and dissolved
oxygen (DO). Microuidic mass sensors and in- situ sensors now allow for continuous measurements.
For example, the Incyte Arc sensors by Hamilton utilize capacitance to measure the polarization of
viable cells. Another example is the Dencytee Total cell density sensors, also by Hamilton, which
rely on optical density or turbidity measurements at near- infrared (NIR) wavelengths.
Measurement of parameters such as pH, pCO2, osmolality, and various metabolites (e.g., glucose,
lactate, glutamine, sodium, potassium) is typically carried out using ofine blood gas analyzers such
as the YSI analyzer and NOVA Biochemical Analyzer.
Productivity estimation is commonly performed through periodic sampling during the fermentation
process. Common quantitation methods include ELISA, Protein A HPLC, and protein biosensor systems
(e.g., ForteBio Octet). AlphaLISA, a high- throughput immunoassay with high sensitivity, reduces testing
time and eliminates some steps involved in traditional ELISA. In the case of bacterial cells, SDS- PAGE
is the most frequently used method for culture analysis and inclusion body (IB) separation.
Glycosylation patterns depend signicantly on cell culture conditions and media/ feed composition, which can have a substantial impact on downstream purication strategies. Glycan mapping
using the UHPLC method is commonly employed for analyzing glycan proles (O- linked and Nlinked). Additionally, high- resolution mass spectrometry aids in determining glycosylation patterns
and degrees (glycoprotein proling). While several orthogonal methods are used for glycan analysis
of the drug substance (Critical Quality Attributes testing), these methods may not be necessary in
the early stages of the manufacturing process.
Both ofine and in- line product monitoring offer several advantages, including the identication
of optimal feeding strategies and culture conditions that can enhance product quality and potentially
reduce the need for additional testing until the nal drug substance preparation (e.g., glycan analysis).
10.7.2 cell haRvest
Turbidity serves as a valuable tool for assessing clarication efciency. It measures the extent of
light scattering when passed through a sample containing suspended particles. This measurement is
expressed as Nephelometric Turbidity Units (NTUs), with lower NTU values indicating better clarication. This method is used to effectively compare different clarication techniques.
The step yield is determined based on the amount of product recovered at the end of the harvest
process. Additionally, for certain processes, estimating the levels of HCP and DNA, predominantly
achieved in downstream purication processes, can offer valuable insights into the achieved level
of purication.
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10.8 UPSTREAM EQUIPMENT
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The technology for upstream unit processes has evolved signicantly over the past few decades.
Originally developed for fermentation, these systems now enable the growth of various cell cultures
capable of yielding a target protein molecule. The biological process of cell growth and expression becomes more sensitive when genetically modied organisms are involved. To address these
challenges, bioreactors are now designed with advanced technology to ensure reproducible batch
yields. This chapter provides an overview of the equipment and operational processes that play a
central role in biopharmaceutical production. While the science of bioprocessing involves a deeper
understanding of the physics and chemistry of bioprocesses, this chapter focuses on the equipment
commonly used in commercial settings.
One of the key criteria for biomanufacturing processes is creating an optimal environment for
all unit operations that support cell growth, isolation/ clarication, and purication to produce the
desired product. Consequently, the facility and equipment are critical components in dening the
process, its performance, and its robustness.
Upstream processing includes cell growth, cell isolation, and harvest, which require equipment
and systems such as:
• Media and Solution Prep Systems.
• Bioreactor Systems for Fermentation.
• Harvest and Clarication Systems.
In addition to the above, other peripheral sanitary equipment essential for the entire
bioproduction process (upstream, downstream, and ll- nish) includes Clean- in- Place (CIP)
systems, sterilization systems, decontamination, biowaste systems, and various disposable/ singleuse systems.
10.9 MEDIA AND SOLUTION PREPARATION SYSTEMS
Vessels and tanks are primarily used for blending, holding bulk liquid volumes, and transport. At
the end of the bioreaction, these vessels are used to introduce materials (media, feed, and other
solutions) into the bioreactor and transfer either parts or the entire reactor volume to the next unit
operation. Stainless steel alloys (particularly 304 and 316) are the most commonly used construction materials for large tanks, but single- use systems are now widely adopted across a broad volume
range. These single- use systems are discussed in more detail in Chapter 9.
These vessels can range from simple storage containers to vessels equipped with overhead
mixers. Features like temperature control, load cells, pH control, liquid/ solid additions, withdrawal,
and gassing can determine the specic use of these tanks or vessels. Jacketed vessels are commonly
used for temperature control, especially in large- scale preparations, where a temperature control
uid circulates inside the jacket to maintain the required temperature in the process uid. Other temperature control components that may be used include an immersion heat exchanger and a heating
blanket.
For vessels with built- in load cells, all additions are based on weight, eliminating the need to
move large tanks for weighing. In large- scale preparations, weight- based solution preparation is
more benecial than volume- based. For lling and transfer, the process length can be established
by setting the overall lling time or dening a lling rate (e.g., kg/ min) for a vessel with a known
volume. pH measurements in bioprocess liquid preparations are typically done ofine through
manual sampling from the preparation tank. However, in high- end systems, such as bioreactor
vessels, continuous online monitoring with a pH probe may be used for pH measurements and
adjustments.
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10.10 BIOREACTOR SYSTEMS
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A bioreactor is a sealed vessel used to support biological systems (e.g., bacterial cells, yeast, mammalian cells) for the production of biopharmaceuticals (such as vaccines and antibodies), primary
and secondary metabolites (including ethanol and biofuel), and even in tissue engineering. The rst
bioreactor was developed in 1857 by Louis Pasteur to study the fundamental principles of fermentation. This milestone in bioprocessing, along with advancements in cell biology and recombinant
DNA technology, paved the way for further developments in bioreactor design and their industrial
applications beyond food products.
Bioreactor design relies on several factors, including cell type, product throughput, quality
control, and mass transfer. The duration required for processes is critical to bioreactor design and
establishing operational parameters. The material used for the bioreactor vessel should endure
repeated sterilization cycles without affecting long- term performance or durability. The preferred
materials are glass (for small- scale benchtop bioreactors) or stainless steel, with 316 SS for wetted
parts and 316L SS for animal cell cultures. Non- product contact surfaces can be made of 304- type
stainless steel. For microbial cultures, a height- to- diameter ratio of 2:1 or 3:1 is preferred for high
gas transfer, while a ratio of 1:1 is common for animal cell cultures. Bioinstrumentation and process
control features to ensure sterility are essential design characteristics for bioreactors. Key functionalities include maintaining sterility, providing aeration (oxygen), removing gaseous products
(carbon dioxide), controlling and optimizing the environment for cell growth (e.g., temperature control, pH, agitation), minimizing evaporation losses, and enhancing process efciency.
10.11 BIOREACTOR TYPES
10.11.1 stiRRed tanK bioReactoR
In bioprocesses, the stirred tank reactor (STR) is the most common and widely used bioreactor type.
They vary in complexity from simple stirred tanks for enzymatic reactions to more advanced aerated
fermenters for metabolic bioconversions. A compressor introduces air from the bottom through
sub- surface spargers to provide necessary aeration. Agitation via an impeller shaft (equipped with
bafes) facilitates mixing and dispersing air bubbles within the vessel’s contents, which are crucial
for bioreactor performance. Stirred tank reactors are highly adaptable, with high kLa (volumetric
mass transfer coefcient) values for gas transfer. Jacketing is often used for heating and cooling.
The height- to- diameter ratio depends on the intended use, ranging from 1 for basic vessels with
limited surface area per unit volume to 3 for large- scale reactors. This type of reactor operates in
one of these modes: batch, fed- batch, or continuous. Various operation modes are explained below
(Figures 10.11 and 10.12).
10.11.2 aiRlift bioReactoR
An airlift or gas- lift bioreactor is a gas- liquid bioreactor that achieves agitation without mechanical
devices, relying on sparged air’s convection. A glass grid aerator aids in humidied air dispersion,
promoting mixing and oxygenation while using less energy than a stirred tank reactor. Only a specic area of the vessel, called the riser, is sparged with gas. Gas holdup and volumetric mass transfer
coefcient (kLa) are inuenced by operating conditions. The medium moves upward in the riser due
to gas holdup and reduced uid density. At the top of the reactor, bubbles disengage, and the denser
medium ows downward into the downcomer, a non- sparged section of the pipe. Oxygen transfer is
often lower than in stirred tank bioreactors due to low shear levels. Airlift bioreactors nd extensive
use in plant and animal cell growth, as well as immobilized biocatalysts.
However, they tend to form dead zones due to non- uniform nutrient distribution and inadequate
mixing (Figure 10.13).
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FIGURE 10.11 Stirred bioreactor
Source: Yassine Mrabet– Own work, CC BY- SA 3.0, https:// comm ons.wikime dia.org/ w/ index.php?curid=
8301 774
FIGURE 10.12 Schematic of an airlift bioreactor. (a) bubble column reactor (BCR), and (b) internal- loop
airlift reactor (ALR)
Source: https:// www.mdpi.com/ 2227- 9717/ 8/ 6/ 713/ htm; http:// crea tive comm ons.org/ licen ses/ by/ 4.0/
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FIGURE 10.13 Fluidized bed bioreactor
Source: YassineMrabet– Own work based on Fluidized Bed Reactor Graphic. JPG (user: Hughesy127), Public
Domain, https:// comm ons.wikime dia.org/ w/ index.php?curid= 8278 465
10.11.3 PacKed bed and fluidized bed bioReactoR
The chemical industry employs packed bed reactors for various processes, such as distillation, absorption, separation, and catalytic reactions. In a packed- bed bioreactor, immobilized
biocatalyst particles (enzyme or microbial cells) ll a tube- shaped vessel. The liquid medium
ows either uphill or downhill through the column, interacting with the catalyst spread along its
length. Rapid liquid phase velocity enhances mass transfer. To improve catalyst conversion, the
process medium is typically recycled through the column multiple times, often requiring an intermediate storage tank. Fluidized beds are a type of packed bed reactor where the liquid medium
ows upwards, allowing the bed to expand at high ow rates, known as extended– or uidized- bed
bioreactors. Biocatalyst particles must have suitable size and density. Continuous movement of
particles prevents channeling and clogging. However, understanding mass and energy transfers
and accounting for potential pressure drops along the reactor’s length are critical challenges in
packed bed reactor design.
Different types of bioreactors include the loop reactor, immobilized cell reactor, solid- phase tray
reactor, rotary drum reactor, agitated tank reactor with a movable impeller, hollow ber reactor, and
the widely popular disposable WAVE™ bioreactor (Figure 10.14).
Several iterations of the Wave bioreactors have been reported in recent patents by Niazi (author
of this book), including a bafed bioreactor (Figures 10.15 and 10.16).
r
−
−Xm XSo/tcYX SM
tc
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FIGURE 10.14 Schematic of a rocking bioreactor
10.12 MODES OF OPERATION
There are four modes of operation available for bioreactors: batch, fed- batch, perfusion, and continuous fermentation.
In the early days of biopharma manufacturing, the batch mode was predominantly used due to
its simplicity. However, over time, fed- batch has become more prominent for microbial and animal
cell culture, offering increased productivity compared to the traditional batch mode. Perfusion, originally considered precise, has also gained renewed interest, especially for animal culture, as an
alternative to fed- batch mode when culture productivity is exceptionally low.
The choice of operation mode by manufacturers is driven by factors such as the type of expression system used, productivity, manufacturing capacity, facility layout, and output.
10.12.1 batch cultuRe
At the start of the process, a batch operation involves a closed vessel system containing all necessary medium components, including the required inoculum for fermentation (cell development).
No subsequent feeding or addition of supplement components is performed, maintaining a constant volume throughout the process. Apart from gases, antifoam, and pH adjusting solutions
added at the beginning, the medium composition must meet all nutritional requirements to
support cell growth and enhance productivity. The substrate concentration remains at its maximum within the reactor, while the cell and product concentration start at their lowest. The entire
batch is harvested, and the fermentation end time is typically determined by product kinetics
(Figure 10.17).
The simplicity of setup and execution led to the widespread application of the batch process,
especially during the initial biotechnology phase involving cell mass production or primary product
production (e.g., vaccine production using microcarriers). A batch process typically includes phases
that dene cell growth: a lag phase, an exponential growth phase, and a stationary phase indicative
of harvesting, followed by preparation for a new batch involving cleaning, sterilizing, and lling;
Figure 10.17).
The total amount of cell mass- produced in each batch process is calculated per the following
equation:
rb = rate of cell mass production per batch cycle and tc = batch cycle time
=
b
=
Equation 10.1
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