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\begin{document}

\title{Adaptive evolution in producing microtiter cultivations generates
genetically stable \emph{Escherichia coli} production hosts for
continuous bioprocessing}



\author[1]{Artur Schuller}%
\author[1]{Monika Cserjan-Puschmann}%
\author[1]{Christoph Köppl}%
\author[1]{Reingard Grabherr}%
\author[2]{Martin Wagenknecht}%
\author[3]{Matteo Schiavinato}%
\author[3]{Juliane Dohm}%
\author[3]{Heinz Himmelbauer}%
\author[1]{Gerald Striedner}%
\affil[1]{Christian Doppler Laboratory for production of next-level biopharmaceuticals in E. coli, University of Natural Resources and Life Sciences}%
\affil[2]{Boehringer Ingelheim RCV GmbH und Co KG}%
\affil[3]{Institute of Computational Biology, University of Natural Resources and Life Sciences}%


\vspace{-1em}



  
  \date{June 15, 2024}


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\selectlanguage{english}
\begin{abstract}
The production of recombinant proteins usually reduces cell fitness and
the growth rate of producing cells. The growth disadvantage favors
faster-growing non-producer mutants. Therefore, continuous bioprocessing
is hardly feasible in \emph{Escherichia coli} due to the high escape
rate. We investigated the stability of \emph{E. coli} expression systems
under long-term production conditions and how metabolic load triggered
by recombinant gene expression influences the characteristics of
mutations. We conducted iterated fed-batch-like microbioreactor
cultivations under production conditions. We used the easy-to-produce
green fluorescent protein (GFP) and a challenging antigen-binding
fragment (Fab) as model proteins, and BL21(DE3) and
BL21\textsuperscript{Q} strains as expression hosts. In comparative
whole genome sequencing analyses, we identified mutations that allowed
cells to grow unhindered despite recombinant protein production. A T7
RNA polymerase expression system is only conditionally suitable for
long-term cultivation under production conditions. Mutations leading to
non-producers occur in either the T7 RNA polymerase gene or the T7
promoter. The host RNA polymerase-based BL21\textsuperscript{Q}
expression system remained stable in the production of GFP in long-term
cultivations. For the production of Fab, mutations in \emph{lacI} of the
BL21\textsuperscript{Q} derivatives had positive effects on long-term
stability. Our results indicate that adaptive evolution carried out with
genome-integrated \emph{E. coli} expression systems in microtiter
cultivations under industrial relevant production conditions is an
efficient strain development tool for production hosts.%
\end{abstract}%



\sloppy


\textbf{1 Introduction}

Fed-batch bioprocessing is the most common cultivation method in
industrial microbial production of biopharmaceuticals. This batch-wise
process essentially includes the repetitive steps of media preparation
and reactor setup, fermentation, and subsequent cleaning in place (CIP)
and sterilization in place (SIP). In terms of time, the actual
fermentation, and particularly the production phase of the recombinant
protein, is relatively short. As a result, continuous production becomes
more and more interesting due to the greatest possible space time yields
and optimal use of the installed assets.\textsuperscript{{[}1{]}}In such
chemostat cultivations, cells are maintained in a steady-state growth
environment by adding fresh medium to the reactor at constant flow.
Simultaneously, the cell suspension, and thus the recombinant protein,
is removed at the same rate.\textsuperscript{{[}2{]}} The growth rate
(\selectlanguage{greek}μ\selectlanguage{english}) can be specified depending on the dilution rate (D). Through this
process, stable volumetric productivity and high space-time yield can be
achieved \textsuperscript{{[}3{]}}. In contrast to fed-batch
fermentation, the average residence time of a producing cell is always
the same, which can be advantageous in terms of product quality.
Examples of microbial continuous processes for the production of
recombinant proteins have already been described in the
literature.\textsuperscript{{[}4-6{]}}

For industrial microbial production of recombinant proteins,
the\emph{Escherichia coli} strain BL21(DE3) is often used due to low
acetate formation and high production rates resulting from the
integrated T7 RNA polymerase (RNAP).\textsuperscript{{[}7, 8{]}} In
combination with a pET series plasmid, which harbors the gene of
interest (GOI) under control of the T7 promoter, extraordinarily high
expression rates can be achieved after induction with the
non-metabolizable lactose analogue isopropyl-D-1-thiogalactopyranoside
(IPTG).\textsuperscript{{[}9{]}}

Genetic heterogeneity caused by metabolic burden and toxicity can be
problematic at all industrial scales, especially for challenging
proteins.\textsuperscript{{[}10-12{]}}In bacterial production processes,
challenging proteins impose adverse effects on host metabolism, even at
low concentrations. Escape variants, which have a growth advantage due
to mutations or plasmid loss, can overgrow high-performing producer
cells, reducing the overall product
yield.\textsuperscript{{[}13-15{]}}Therefore, longer production phases,
or even continuous production mode, are hardly feasible in such \emph{E.
coli} expression systems.

To obtain stable, high-yield, and predictable \emph{E. coli} production
hosts, engineered producer strains must focus on reducing metabolic load
and genetic escape. Attempts to reduce process instability caused by
metabolic burden have been made on both the genetic and bioprocessing
levels. The metabolic burden and increased selection pressure can be
reduced by decoupling growth and production in cascading chemostat
cultivations with two bioreactors.\textsuperscript{{[}6{]}} Genetic
escape can be reduced by removing insertion elements (IS), and by
deleting \emph{recA} or error-prone DNA polymerase
genes.\textsuperscript{{[}16{]}} Another promising strategy uses the
directed evolution approach combined with fluorescence-activated cell
sorting to select cells with a lower plasmid mutation
rate.\textsuperscript{{[}17{]}}

In previous studies, we showed that genomic integration of the GOI under
the control of the strong T7 promoter reduces the metabolic burden
because plasmid-mediated metabolic load is eliminated, and strong
expression has been shown from even a single copy of the GOI. However,
after approximately seven doublings under production conditions,
mutations in the T7RNAP gene lead to a faster growing non-producing cell
population. This phenomenon can be excluded in systems using the host
RNAP-specific A1 promoter because full functionality of the host RNAP is
required for cell growth.\textsuperscript{{[}18{]}}

In combination with a directed evolution approach, this would allow
characterization of mutated production hosts, which have adapted
themselves to the metabolic load triggered by recombinant protein
production and could potentially enable continuous protein production.
Moreover, a directed evolution approach can circumvent the complexity of
the process of biogenesis and its adverse effects on the host
cell.\textsuperscript{{[}19-21{]}}

For example, in the study by Walker et al. \textsuperscript{{[}22{]}},
derivatives of BL21(DE3) were adapted by directed evolution to produce
cell membrane proteins that are toxic to host cells. These strains are
currently widely used for production of a variety of membrane proteins
and toxic proteins.

In the present study, we performed iterated carbon-limited
fed-batch-like microbioreactor cultivations under production conditions.
The goal was to investigate whether and how long-term metabolic load
triggered by the production of recombinant proteins influences the
characteristics of mutations occurring in different
genome-integrated\emph{E. coli} production
systems.\textsuperscript{{[}23, 24{]}} We compared the host
RNAP-dependent BL21\textsuperscript{Q} A1 expression system
(BQ\textless{}A1\textgreater{}) \textsuperscript{{[}18{]}} with the
T7-based BL21(DE3) expression system (B3\textless{}T7\textgreater{}). To
study mutation characteristics based on the protein of interest (POI),
we used the easy-to-produce protein GFPmut3.1 and the challenging
protein Fab fragment dFTN2 as model proteins. To investigate the
underlying mutations in the different clones in more detail, we
performed comparative whole genome sequencing analyses. We also
performed long-term chemostat cultivations in lab-scale bioreactors with
the above-mentioned clones and additional robust production strains
obtained by the directed evolution approach.

\textbf{2 Materials and methods}

\textbf{2.1 Construction of expression systems}

\textbf{2.1.1 Strains}

\emph{E. coli} K-12 NEB5-\selectlanguage{greek}α\selectlanguage{english} {[}\selectlanguage{greek}\emph{fhuA2Δ(argF-lacZ)U169 phoA gln V44
Φ80 Δ(lacZ)M15 gyrA96 recA1 relA1 endA1 thi-1 hsdR17}\selectlanguage{english} {]} was obtained
from New England Biolabs (NEB, Ipswich, USA) and used for all cloning
procedures. For recombinant protein expression, linear DNA cartridges
controlled by the T7 promoter were integrated into the bacterial
chromosome at the attTn7 site of \emph{E. coli} BL21(DE3) {[}\selectlanguage{greek}\emph{fhuA2
{[}lon{]} ompT gal (λ DE3) {[}dcm{]} [?]hsdS λ DE3 = λ sBamHIo [?]EcoRI-B
int::(lacI::PlacUV5::T7~gene1) i21 [?]nin5} {]} (NEB). Hereafter, this
strain is referred to as B3\textless{}T7-\emph{gene} \textgreater{}.
Linear DNA cartridges controlled by the A1 promoter were integrated into
the bacterial chromosome at the attTn7 site of \emph{E. coli} BL21
{[}fhuA2 {[}lon{]} ompT gal {[}dcm{]} \selectlanguage{greek}Δ\selectlanguage{english}hsdS{]} (NEB) containing
the\emph{lacI\textsuperscript{Q}} promoter as described by Glascock and
Weickert.\textsuperscript{{[}25{]}} Briefly, the pETAmp-lacIq plasmid
was constructed for integration of the
\emph{lacI\textsuperscript{Q}}promoter in \emph{E. coli} BL21. This
plasmid contains the ampicillin resistance gene flanked by FRT sites and
the \emph{lacI} gene controlled by the \emph{lacI\textsuperscript{Q}}
promoter. The pBR322 ori and\emph{lacI} were amplified from pET30a using
the overhang PCR technique to introduce a C -\textgreater{} T mutation
in the \emph{lacI} promoter. The linear \emph{lacI\textsuperscript{Q}}
DNA cartridge for genome integration was amplified using
Q5\selectlanguage{ngerman}\textsuperscript{®} High-Fidelity DNA Polymerase (NEB) according to
the manufacturer's manual. Integration into the bacterial chromosome
occurred at the lac operon site of\emph{E. coli} BL21 carrying the pSIM5
plasmid.\textsuperscript{{[}26{]}} This strain was designated as
BL21\textsuperscript{Q} and referred to hereafter as
BQ~\textless{}A1-\emph{gene} \textgreater{}.

The cytosolic protein GFPmut3.1 was used as a recombinant
``easy-to-produce'' model protein.\textsuperscript{{[}27{]}} An
antigen-binding fragment (Fab) against TNF-\selectlanguage{greek}α \selectlanguage{english}(FTN2) with the DsbA signal
sequence (dFTN2) was used as a recombinant ``challenging'' model
protein.\textsuperscript{{[}28{]}}

\textbf{2.1.2 Plasmids and integration of expression cassettes into
the\emph{E. coli} chromosome}

Plasmids harboring the integration cassettes of either GFPmut3.1 or
dFTN2 were constructed and integrated into the \emph{E. coli} chromosome
as described by Fink et al..\textsuperscript{{[}28{]}}

\textbf{2.2 Microbioreactor cultivations}

The strains were cultivated in the BioLector\selectlanguage{ngerman}® microfermentation system
in 48-well Flowerplates® (m2p-labs, Baesweiler, Germany) as described by
Török et al..\textsuperscript{{[}29{]}} Synthetic feed in time (FIT)
fed-batch medium containing 1 g/L glucose and 16.5 g/L dextran as carbon
sources (m2p-labs GmbH, Baesweiler, Germany) was used with the following
additions (g/L): 27.40 MOPS, 6.54
(NH\textsubscript{4})\textsubscript{2}SO\textsubscript{4}, 1.96
K\textsubscript{2}HPO\textsubscript{4}, 1.96 trisodium
citrate·2H\textsubscript{2}O, 1.31
Na\textsubscript{2}SO\textsubscript{4}, 0.65 NH\textsubscript{4}Cl, 0.33
MgSO\textsubscript{4}·7H\textsubscript{2}O, and 0.0065 Thiamin·HCl.

The trace element solution contained (mg/L): 0.36
ZnSO\textsubscript{4}·7H\textsubscript{2}O, 0.33
CuSO\textsubscript{4}·5H\textsubscript{2}0, 0.20
MnSO\textsubscript{4}·H\textsubscript{2}O, 27.30
FeCl\textsubscript{3}·6H\textsubscript{2}O, 21.84 Titriplex III, 0.36
CoCl\textsubscript{2}·6H\textsubscript{2}O, and 1.31
CaCl\textsubscript{2}·2H\textsubscript{2}O. Immediately prior to
inoculation, 0.6\% (v/v) of the glucose releasing enzyme mix (EnzMix)
was added. The GFPmut3.1 expression level was monitored at an excitation
of 488 nm and emission of 520 nm. The signal is given in relative
fluorescence units (rfu). The cell dry matter (CDM, given in g/L) was
calculated from light scatter signals using calibration settings
obtained by linear regression analysis. The cycle time for all
parameters was 20 min.

The initial cell density was equivalent to an optical density of
OD\textsubscript{600} = 0.3. For inoculation of passage 1, a deep-frozen
(-80°C) working cell bank (WCB; OD\textsubscript{600} = 3.5) was thawed
and biomass harvested by centrifugation (7500 rpm, 5 min). Cells were
washed with 500~\selectlanguage{greek}μ\selectlanguage{english}L of the corresponding medium to remove residual
glycerol and centrifuged. Pellets were then re-suspended in the total
cultivation medium. All subsequent passages were inoculated with induced
cells from the previous passage, but without adding batch glucose again,
keeping the cells in carbon-limited conditions. All cultivations were
performed at 30\selectlanguage{ngerman}°C. Recombinant gene expression was induced with IPTG at
a final concentration of 0.5 mM.

\textbf{2.3 Fed-batch cultivations}

For fed-batch fermentation, cells were grown in a 1.5 L (1.2 L working
volume, 0.4 L minimal volume) DASGIP\textsuperscript{®} Parallel
Bioreactor System (Eppendorf AG, Hamburg, DE) equipped with standard
control units. The pH was maintained at 7.0 ± 0.05 by the addition of
12.5\% ammonia solution (Thermo Fisher Scientific, Waltham, USA). The
temperature was maintained at 37 ± 0.5°C during the batch phase and
decreased to 30 ± 0.5°C during the feed phase. The dissolved oxygen
(O\textsubscript{2}) level was stabilized at \textgreater{} 30\%
saturation by controlling the stirrer speed and aeration rate. Foaming
was suppressed by the addition of antifoam suspension (Glanapon, 2000,
Bussetti, AT). For inoculation, a deep-frozen (-80°C) WCB vial was
thawed and 1 mL transferred aseptically to a 250 mL pre-culture shake
flask containing 25 mL M9ZB \textsuperscript{{[}30{]}} for cultivation
for at least 8 hours at 37°C. Subsequently, a volume equivalent to 25
OD\textsubscript{600} units (25/OD\textsubscript{600} = volume in mL)
was transferred aseptically to the bioreactors.

Feeding was initiated when the culture, grown to 10 g/L CDM in 0.6 L
batch medium, entered the stationary phase. A fed-batch regimen with
exponential carbon-limited substrate feed was used to provide a constant
growth rate of \selectlanguage{greek}µ \selectlanguage{english}= 0.1/h over 19 hours or 2.74 doublings. The substrate
feed was controlled by increasing the pump speed according to the
exponential growth algorithm, \selectlanguage{greek}\(x=x_{0}e^{\text{µt}}\selectlanguage{english}\), with superimposed
feedback control of weight loss in the substrate bottle. The CDM yield
coefficient on glucose was 0.33 g/g. Therefore, the feed medium provided
66 g/L glucose and sufficient components to yield a final CDM
concentration of 30 g/L in 1.2 L. The expression system was induced by
adding IPTG to the reactor to yield a concentration of 10 \selectlanguage{greek}µ\selectlanguage{english}mol/g CDM.
The minimal medium was prepared as previously
described.\textsuperscript{{[}15{]}}

\textbf{2.4 Chemostat cultivations}

Chemostat cultivations were run at a density of 30 g/L CDM and a
dilution rate of D = 0.1/h in a working volume of 0.67 L. The starting
batch process was followed by a fed-batch phase. The batch volume was
set to 400 mL and the batch medium allowed production of 4.72 g CDM; the
feed volume was set to 270 mL with medium designed to produce another
15.4 g of CDM, which corresponds to a final CDM concentration of 30 g/L.
In the next step, the bioreactor was shifted to chemostat mode with a
dilution rate of 67 mL/h. The medium was used as continuous feed to the
reactor to provide nutrients to maintain a CDM concentration of 30 g/L.
The temperature was decreased to 30\selectlanguage{ngerman}°C and recombinant protein production
induced with 10 \selectlanguage{greek}µ\selectlanguage{english}mol IPTG/g CDM. Volume was kept constant via an
immersion tube adjusted to the right height of the liquid surface,
ensuring that the bleed pump was working at a higher rate than the feed
pump.

\textbf{2.5 Analysis of recombinant protein expression}

\textbf{2.5.1 Flow cytometry}

A CytoFLEX S flow cytometer (Beckman Coulter, Brea, USA) was used to
monitor subpopulations of GFPmut3.1-producing cells. Cells were diluted
1/2025 in PBS. Excitation of GFPmut3.1 fluorescence was performed at 488
nm, with subsequent emission measured using the FL1 Channel (525BP40-A).
Data were recorded for 15,000 cells per sample at \textasciitilde{}300
events/s and were analyzed by Kaluza analysis software (Beckman
Coulter).

\textbf{2.5.2 Off-line measurement of GFP fluorescence}

Off-line fluorescence measurements were made using a Tecan analyzer
infinite 200Pro. For GFPmut3.1, the excitation wavelength was 485 nm and
the emission wavelength 520 nm. Calibration with the in-house purified
target protein was used for quantification.

\textbf{2.5.3 Cell lysis}

Samples from microbioreactor experiments were taken at the end of each
passage. Samples from fed-batch cultivations were taken every 2 hours
and samples from chemostat cultivations every 24 hours. The sample
volume for intracellular protein quantification was calculated as
3.5/OD\textsubscript{600} = volume in mL, which corresponds to
\textasciitilde{}1 mg CDM. Cell lysis for quantification of
intracellular recombinant proteins was performed as described
elsewhere.\textsuperscript{{[}28{]}}

\textbf{2.5.4 Quantification of soluble recombinant protein}

Recombinant GFPmut3.1 and FTN2 were quantified by ELISA as described
previously.\textsuperscript{{[}28, 31{]}}

\textbf{2.5.5 Fab expression pattern on dot blot}

The FTN2 samples from microbioreactor cultivations were analyzed
initially in a high-throughput manner using dot blot to obtain a simple
yes-or-no-answer. For that purpose, 200 \selectlanguage{greek}µ\selectlanguage{english}L of cell suspension were
centrifuged in a 48-deep-well plate at 1200 rpm for 10 min. The
supernatant was removed and pelleted cells resuspended in 400 \selectlanguage{greek}µ\selectlanguage{english}L TE
buffer (pH 8). Resuspended cells were incubated for 1 hour at 60\selectlanguage{ngerman}°C with
shaking, and then subsequently centrifuged as described above. The
supernatant (3 \selectlanguage{greek}µ\selectlanguage{english}L) was pipetted on a nitrocellulose membrane. Blocking,
incubation with antibodies, and membrane development were performed as
described below for Western blotting.

\textbf{2.5.6 Fab expression pattern via Western blot}

Soluble Fab expression, IB formation, and basal expression levels were
analyzed by Western blot as described
previously.\textsuperscript{{[}28{]}}

\textbf{2.5.7 Paired-end library preparation and whole genome
sequencing}

Genomic DNA extraction was achieved by standard phenol:chloroform
extraction as described previously.\textsuperscript{{[}32{]}} The
quality of the genomic DNA was checked on a 0.6\% standard agarose gel
stained with ethidium bromide using a NanoDrop ND-1000 spectrophotometer
(ThermoFisher, Waltham, MA, USA). Quantification was performed using a
Qubit 3.0 Fluorometer and a dsDNA BR assay kit (ThermoFisher, Waltham,
MA, USA). Paired-end libraries for Illumina sequencing were prepared
using a TruSeq Nano DNA Low Throughput Library Prep kit (Cat \#
20015964, Illumina, San Diego, CA, USA) according to the supplier's
instructions. Briefly, library construction began with fragmentation of
200 ng of genomic DNA to a peak fragment size of 550 bp using a Covaris
M220 instrument (Covaris, Woburn, MA, USA) with the following settings:
45 s at 20\% duty cycle, intensity 50, temperature 20\selectlanguage{ngerman}°C, and 200 cycles
per burst. The DNA fragments were then purified using SPB beads included
in the TruSeq kit, followed by end-repair, A-tailing, and ligation of
Illumina adapters to the ends of the fragments. After SPB purification,
the library was PCR-amplified using the following cycling conditions:
initial denaturation at 95°C for 3 min, followed by 8 cycles at 98°C for
20 s, 60°C for 15 s, and 72°C for 30 s, and a final extension at 72°C
for 5 min. After PCR clean-up, 1 \selectlanguage{greek}µ\selectlanguage{english}L of the library was used for
validation in a 2100 Bioanalyzer (Agilent, Santa Clara, CA, USA).
Library quantification was accomplished on a Qubit 3.0 Fluorometer using
a dsDNA BR assay kit. The library was then sequenced at the VBCF NGS
Core facility (Vienna, Austria) on an Illumina HiSeq 2500 sequencing
instrument using v4 sequencing chemistry and a 2\selectlanguage{ngerman}×125 nt paired-end
sequencing protocol.

Raw genomic short reads from all wild-type and mutant strains were
quality trimmed with Trimmomatic
v0.35.\textsuperscript{{[}33{]}}(ILLUMINACLIP:TruSeq3-PE.fa:2:30:10
LEADING:20 TRAILING:20 SLIDINGWINDOW:4:20 AVGQUAL:20 MINLEN:50). We
generated four reference sequences that represented our versions of the
T7 and A1 wild-type strains: B3\textless{}T7-GFP\textgreater{} wt,
B3\textless{}T7-Fab\textgreater{} wt, BQ\textless{}A1-GFP\textgreater{}
wt, and BQ\textless{}A1-Fab\textgreater{} wt. Variants between the T7
wild-type strains and the publicly available BL21(DE3) genome (NCBI
Reference Sequence: NC\_012892.2) were extracted using their
quality-trimmed reads. The same was repeated for A1 wild-type strains
using the BL21 genome.\textsuperscript{{[}34{]}} To extract the
variants, we used breseq \textsuperscript{{[}35{]}} (limit fold-coverage
150, minimum mapping quality 20, maximum read mismatches 15, no junction
prediction, require match-fraction 0.5). The four updated references
were obtained introducing the found variants with gdtools APPLY provided
with breseq.

Using the quality-trimmed reads of the mutant strains, one genome
sequence was assembled for each mutant. The peak insert size of each
sequencing library was determined by mapping up to 10,000 read pairs
with HISAT2 \textsuperscript{{[}36{]}} on the appropriate reference
genome sequence. Quality-trimmed reads were then down-sampled to a
coverage of 150× using seqtk sample (https://github.com/lh3/seqtk).
Down-sampled reads were assembled using
SOAP-denovo\textsuperscript{{[}37{]}} (-M 3 -L 100 -K 75 -k 25 -d 5 -D
5). The peak insert size determined for each library was used to
calibrate the genome assembler. The specified config file parameters
were: reverse\_seq=0, asm\_flags=3, max\_rd\_len=125,
rd\_len\_cutoff=125, rank=1, map\_len=30. Assembly metrics (contig N50
length and contig N90 length) were assessed using
Biopython.\textsuperscript{{[}38{]}}

Single nucleotide polymorphisms (SNPs) and short insertion/deletion
(indel) variants up to 20 bp in length between mutant and wild-type
strains were extracted. Short reads from each mutant were mapped onto
their corresponding wild-type reference using
breseq.\textsuperscript{{[}35{]}} The program was run in polymorphism
mode (polymorphism prediction, polymorphism frequency cutoff 0.05) using
the same parameters as for the generation of mutated references. True
variants were then selected by mapping the contigs of the assembled
genomes of the mutants; only variants with both read and contig mapping
were retained. The clonality of each variant was inferred from the 'AF'
field in the VCF files.

The presence of candidate genome rearrangements was screened using the
assembled genome sequences for the mutants. Contigs were mapped onto the
corresponding wild-type reference sequence with
nucmer\textsuperscript{{[}39{]}} (mum, breaklen 10, mincluster 500,
delta, diagfactor 0.05, maxgap 30000, minmatch 50). The produced mapping
records were then passed to the show-diff tool, available with nucmer,
which highlights potential rearrangements and their type.



\textbf{3 Results and Discussion}

\textbf{3.1 Directed evolution and isolation of protein production
strains}

We applied repeated fed-batch-like cultivations in a 48-well
microbioreactor system in order to characterize mutation patterns
triggered by recombinant protein production under long-term
cultivations. Twenty-four wells were inoculated with the host
RNAP-dependent A1 expression system (BQ\textless{}A1\textgreater{}) and
24 with the T7 RNAP-dependent BL21(DE3) expression system
(B3\textless{}T7\textgreater{}), in which both strains have the GOI
integrated into the chromosome. GFP-producing cells were induced with 10
\selectlanguage{greek}µ\selectlanguage{english}mol IPTG/g CDM at the beginning of each passage, which corresponds to
full induction of recombinant protein production. Fab-producing cells
were induced after 6 hours of the first passage and cultivated in medium
that already contained 10 \selectlanguage{greek}µ\selectlanguage{english}mol IPTG/g CDM in all other passages. Cells
were passaged several times until no difference in growth behavior was
observed. We performed the experiments with two different model
proteins, the easy-to-produce protein GFPmut3.1 and challenging protein
dFTN2. For GFP-producing strains, the fluorescence of GFP was used to
distinguish between producing and non-producing clones over a period of
six passages and a total of 42 generations. Fab-producing cells were
cultivated over a period of three passages (21 generations) and dot blot
analysis performed to detect producing clones.

\textbf{3.1.1 Evolution of GFP-producing \emph{E. coli} strains}

Induction of cells producing GFP under the control of the A1 promoter,
BQ\textless{}A1-GFP\textgreater{}, showed no reduction in cell growth
(Figure S1). The cells were able to reach the calculated end biomass of
6 g/L CDM in each of the six passages. Similarly, the productivity
remained unchanged and was always approximately 25 rfu/g CDM at the end
of each passage (Figure S2). Flow cytometry revealed continuous
homogeneous populations of all 24 measured cultivations, as depicted for
clones \#E1 and \#E2 in Figure 1B.

Unlike cells producing GFP under the control of the strong T7 promoter
(B3\textless{}T7-GFP\textgreater{}), induction of the GOI at the
beginning of the cultivation led to a considerable reduction in cell
growth. In passage 1, cells reached a final biomass of only 1.5 g/L CDM.
However, all 24 T7 strains were able to reach the calculated end biomass
of 6 g/L already in passage 2. This was accompanied by a strong
reduction in productivity. As shown in Figure 1A, the first
subpopulations of non-producers and weak producers appeared in passage 2
and continued to outgrow producing strains over the period of six
passages, as shown by derivatives \#F6 and \#F7. However, in derivatives
\#B8, \#E7, and \#F6, we found mixed populations of non-producers, weak
producers, and strong producing cells, which could be maintained over
the whole period of six passages.

To separate single-cell colonies from this mixture, we streaked the
cells on agar plates after passage 6 and analyzed the isolated colonies
by flow cytometry. We isolated 10 homogeneous subpopulations from weak
producing (subclone 1, S1) to strong producing (subclone 10, S10) cells
(Figure 1C).\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image1/image1}

\end{center}
\end{figure}

\textbf{Figure 1.} Analysis of single cell expression using GFP as a
model protein. (A-B) Flow cytometry over the course of six passages. (A)
Derivatives of B3\textless{}T7-GFP\textgreater{}. (B) Derivatives of
BQ\textless{}A1-GFP\textgreater{}. (C) Derivatives of
B3\textless{}T7-GFP\textgreater{} separated and categorized according to
the strength of their expression. S1: weak producing to S10: strong
producing subclones.

\textbf{3.1.2 Evolution of Fab-producing \emph{E. coli} strains}

Fab-producing strains were cultivated in the same way as described for
the GFP-producing strains, except cells were kept induced for a total of
three passages. All Fab-producing clones exhibited reduced growth rates
in passage 2 but were able to recover and exhibited unrestricted growth
in passage 3 (Figure 2A). Dot blot analysis revealed that none of the 24
T7 clones produced Fab anymore, whereas three A1 clones exhibited the
desired properties of fast cell growth and an ability to produce the
challenging protein. To ensure homogeneous populations, the three A1
clones were separated on agar plates. Four colonies were picked from
each plate to inoculate further cultivation in media containing IPTG to
confirm their productivity. As shown in Figure 2B and C, we were able to
isolate three subclones (\#B4.1, \#D2.4, and \#E2.1) that produced Fab
without reducing cell growth, but in different quantities; clone \#E2.1
produced comparable amounts of Fab as the non-mutated
BQ\textless{}A1-Fab\textgreater{} wt strain, whereas the other two
subclones exhibited reduced productivity.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image2/image2}

\end{center}
\end{figure}

\textbf{Figure 2.} Growth characteristics and product formation of
Fab-producing strains. (A) Growth behavior of
BQ\textless{}A1-Fab\textgreater{} (red lines) and
B3\textless{}T7-Fab\textgreater{} (blue lines) over the course of three
passages. The dashed vertical line indicates the time point of
induction. (B) Cells were fractionated on agar plates and re-tested for
growth and productivity. (C) The specific concentration of Fab is given
in milligrams Fab per gram of cell dry matter {[}mg/g CDM{]}.

\textbf{3.2 Comparative genomic analysis of \emph{E.
coli}BQ\textless{}A1\textgreater{} and B3\textless{}T7\textgreater{}
derivatives}

To search for genetic factors involved in the ability to maintain a high
growth rate under production conditions, we sequenced the genomes of 11
B3\textless{}T7-GFP\textgreater{} and 2
BQ\textless{}A1-GFP\textgreater{} derivatives (Table 1) and 3
B3\textless{}T7-Fab\textgreater{} and 4
BQ\textless{}A1-Fab\textgreater{} derivatives (Table 2). We found one to
three mutations (SNPs or indels) per genome, corresponding to the
mutation rate for \emph{E. coli} of approximately 1 \selectlanguage{ngerman}×
10\textsuperscript{-3} per generation for cells cultured under standard
laboratory conditions.\textsuperscript{{[}40{]}} Notably, the
spontaneous mutation rate is stress-dependent
\textsuperscript{{[}41{]}}and mutants that result in an increased growth
rate under production conditions are more likely to be selected. We
applied read mapping and a\emph{de novo} assembly approach to examine
SNPs and short indels, as well as detect larger deletions or
subpopulations of genomes.

\textbf{3.2.1 Genetic escape variants of GFP-producing derivatives}

Out of the 24 BQ\textless{}A1-GFP\textgreater{} derivatives isolated
after directed evolution, the genomes of two randomly selected
derivatives were sequenced, since all of them exhibited the same
phenotype after six passages. Interestingly, both derivatives exhibited
exactly the same mutation within the
\emph{lacI\textsuperscript{Q}}promoter (Table 1). Next to the introduced
C\selectlanguage{english}-T substitution\textsuperscript{{[}25{]}} within the -35 region of the
\emph{LacI}promoter, we found a G\selectlanguage{english}-A transition. However, this mutation
has no influence on the basal expression or tunability of the A1
expression system (Figure S3).

Of the 24 B3\textless{}T7-GFP\textgreater{} derivatives, we sequenced
the genomes of the 11 isolated subclones (Figure 1C). The mutations
found in the B3\textless{}T7-GFP\textgreater{} derivatives can be
divided into three clusters: mutations within the T7 promoter or T7 RNAP
gene, mutations within genes involved in the metabolism and transport of
various sugars, and mutations within the \emph{lac} operon. Among the
non-producing and weakly producing strains (subclones S1-S4, Figure 1C),
mutations within the T7 RNAP gene or the T7 promoter were responsible
for the observed phenotype. Derivative \#E7P6 S1 had a 13-bp deletion
within the T7 promoter encompassing almost the complete T7 promoter
sequence. In \#A5P6 S1, 10,872 bp of the DE3 lysogen were deleted,
harboring the complete sequence of the T7 RNAP gene. In general, no
mutational hotspot within the T7 RNAP gene was seen. In each sequence of
the B3\textless{}T7-GFP\textgreater{} derivatives, which had a mutation
in the T7 RNAP, we found a different SNP or indel that caused a loss of
function of the T7 RNAP.

As a second cluster, we identified mutations within genes involved in
the metabolism and transport of various sugars. One mutation found in
the A1 and T7 expression system was a C\selectlanguage{english}-T mutation within the\emph{gntT}
operator site.\textsuperscript{{[}42, 43{]}} The influence of the
gluconate transporter on lactose-inducible gene regulation and its
influence on growth during production cannot be fully explained because
the medium used for cultivation did not contain any gluconate.
Derivative \#B8P6 S2 exhibits a T \selectlanguage{english}- G transversion within the promoter
sequence of \emph{setA} . SetA is an efflux pump capable of transporting
a range of sugars and sugar analogues. Cells overexpressing SetA exhibit
decreased accumulation of lactose and IPTG.\textsuperscript{{[}44{]}} We
speculate that this mutation increases the SetA expression rate,
preventing intracellular accumulation of IPTG and reducing the induction
of the GOI.

Derivative \#A5P6 S5 has a T\selectlanguage{english}-C mutation within \emph{ptsI} , leading to
a L256P substitution. PtsI is a cytoplasmic protein that serves as the
gateway for the phosphoenolpyruvate:sugar phosphotransferase system
(PTS\textsuperscript{sugar}). \emph{E. coli} mutants with \textless{}
1\% residual PtsI activity are unable to consume the PTS sugars glucose,
fructose, mannose, mannitol, sorbitol, N-acetylglucosamine, and
N-acetylmannosamine, and the non-PTS sugars glycerol, melibiose,
maltose, and lactose \textsuperscript{{[}45{]}}. The L256P substitution
may reduce PtsI activity to a level at which glucose can still be used
as a substrate, but the intake of IPTG is reduced, which also reduces
induction of the GOI.

In addition to a mutation within the T7 RNAP gene, derivative \#A5P6 S7
also exhibited a mutation within \emph{malP} . The
(T)\selectlanguage{english}\textsubscript{5?4}mutation causes a frameshift and probably complete
loss of function. MalP is involved in maltose metabolism and has high
affinity for short, linear \selectlanguage{greek}α-\selectlanguage{english}1,4 linked
oligoglucosides.\textsuperscript{{[}46{]}} A mutation within \emph{srlR}
was found in derivative \#B8P6 S8. This gene encodes the glucitol
repressor GutR and is a DNA-binding transcription factor that represses
the \emph{gut} operon involved in the transport and utilization of
glucitol.\textsuperscript{{[}47{]}}

A mutation in the type III CoA transferase gene,
\emph{caiB}\textsuperscript{{[}48{]}}, was found in four derivatives. It
is unlikely that the mutation in \emph{caiB} influences recombinant
protein production or growth behavior, because all B8 derivatives, which
can be classified as both strong and weak producers, exhibit this
mutation.

Mutations in the third cluster comprise the \emph{lac} operon itself.
Two mutations were found in the pLac promoter in derivatives belonging
to strongly producing subclones S9 and S10. The positive properties of
mutations in pLac on \emph{lac} -regulated recombinant gene expression
are already known \textsuperscript{{[}19{]}}; expression of
the\emph{lac} operon is reduced, including reduced expression of LacY,
the sugar transport protein. Consequently, less IPTG enters the cell,
which weakens induction of the GOI.\textsuperscript{{[}49{]}} The
metabolic burden is reduced, explaining why the selected derivatives no
longer have a disadvantage in growth. We speculate that this phenomenon
also applies to the mutations observed in other genes, such as
\emph{setA} ,\emph{ptsI} , \emph{malP} , \emph{srlR,} and \emph{gntT,}
which are involved in the metabolism of various sugars, even though they
are not known to be directly related to IPTG or glucose transport into
the cell. However, a negative influence on recombinant protein
production can only be clearly attributed in the case of \emph{ptsI} ,
as this mutation does not occur in combination with a mutation in the T7
promoter or in the T7 RNAP gene. Unfortunately, this is not the case
with mutations in\emph{setA} and \emph{malP} . However, the mutations in
\emph{srlR} and\emph{gntT} clearly have a positive influence on the
growth rate under conditions of recombinant protein production.

\textbf{Table 1.} Mutations in GFP-producing derivatives of
B3\textless{}T7-GFP\textgreater{} and BQ\textless{}A1-GFP\textgreater{}
isolated after fractionation.\selectlanguage{english}
\begin{longtable}[]{@{}lllllll@{}}
\toprule
\textbf{Strain} & \textbf{Clone} & \textbf{Position} & \textbf{Mutation}
& \textbf{Region} & \textbf{Gene} & \textbf{Function}\tabularnewline
\midrule
\endhead
B3\textless{}T7-GFP\textgreater{} & E7P6 S1 & 3800855 & \selectlanguage{greek}Δ \selectlanguage{english}13 bp &
Intergenic & -73 from \emph{gfpmut3.1} & T7 promoter\tabularnewline
& A5P6 S1 & 743064 & \selectlanguage{greek}Δ \selectlanguage{english}10,872 bp & Coding & - & Part of \selectlanguage{greek}λ
\selectlanguage{english}DE3\tabularnewline
& B8P6 S2 & 43017 & GGT - GTT (G98V) & Coding & \emph{caiB} & Type III
coA transferase\tabularnewline
& & 80353 & T - G & Intergenic & -73 from \emph{setA} & Sugar
transporter\tabularnewline
& & 3800868 & +TA & Intergenic & -72 from \emph{gfpmut3.1} & T7
promoter\tabularnewline
& A5P6 S3 & 752558 & GGA - TGA (G716*) & Coding & T7 RNAP gene &
Enterobacteria phage T7 RNAP\tabularnewline
& A5P6 S4 & 753056 & TTC - CTC (F882L) & Coding & T7 RNAP gene &
Enterobacteria phage T7 RNAP\tabularnewline
& A5P6 S5 & 2407211 & CTG - CCG (L256P) & Coding & \emph{ptsI} &
Bacterial phosphotransferase system\tabularnewline
& B8P6 S6 & 43017 & GGT - GTT (G98V) & Coding & \emph{caiB} & Type III
coA transferase\tabularnewline
& & 3800859 & +C & Intergenic & -81 from \emph{gfpmut3.1} & T7
promoter\tabularnewline
& A5P6 S7 & 751679 & CGC - TGC (R423C) & Coding & T7 RNAP gene &
Enterobacteria phage T7 RNAP\tabularnewline
& & 3412013 & (T)\textsubscript{5 - 4} & Coding & \emph{malP} &
Maltodextrin phosphorylase monomer\tabularnewline
& B8P6 S8 & 43017 & GGT - GTT (G98V) & Coding & \emph{caiB} & Type III
coA transferase\tabularnewline
& & 2669647 & \selectlanguage{greek}Δ \selectlanguage{english}2 bp & Coding & \emph{srlR} & GutR transcriptional
repressor\tabularnewline
& B8P6 S9 & 43017 & GGT - GTT (G98V) & Coding & \emph{caiB} & Type III
coA transferase\tabularnewline
& & 335936 & C - T & Intergenic & -104 from \emph{lacZ} & Upstream
region of pLac\tabularnewline
& & 3406745 & C - T & Intergenic & -168 from \emph{gntT} & \emph{gntT}
operator site\tabularnewline
& E7P7 S10 & 335936 & C - T & Intergenic & -104 from \emph{lacZ} &
Upstream region of \emph{pLac}\tabularnewline
BQ\textless{}A1-GFP\textgreater{} & E1P6 & 337099 & G - A & Intergenic &
-186 from \emph{lacI} & \emph{pLacI\textsuperscript{Q}}\tabularnewline
& & 3375909 & C - T & Intergenic & -168 from \emph{gntT} & \emph{gntT}
operator site\tabularnewline
& E2P6 & 337099 & G - A & Intergenic & -186 from \emph{lacI} &
\emph{pLacI\textsuperscript{Q}}\tabularnewline
& & 3375909 & C - T & Intergenic & -168 from \emph{gntT} & \emph{gntT}
operator site\tabularnewline
\bottomrule
\end{longtable}

\textbf{3.2.2 Genetic escape variants of Fab-producing derivatives}

In the case of Fab-producing clones, we found only one mutation per
sequenced genome, which is attributable to the shorter cultivation time
and lower generation number.\textsuperscript{{[}40{]}} In addition, the
production of a challenging protein exerts high selection pressure,
which means that single mutations leading to a faster-growing population
are selected much faster. As mentioned above, we could not find a
B3\textless{}T7-Fab\textgreater{} derivative that was able to produce
Fab. The three sequenced genomes exemplified that this common phenotype
was always due to a mutation in the T7 RNAP gene (Table 2). In contrast,
we were able to isolate three BQ\textless{}A1-Fab\textgreater{}
derivatives capable of producing Fab without limiting growth. In
addition, we sequenced the BQ\textless{}A1-Fab\textgreater{}
non-producer derivative C3. In the latter case, the reason for the lack
of production is a mutation in the -35 region of the A1
promoter.\emph{In vivo,} the promoter function correlates with the
degree of sequence homology of -35 and -10 with the consensus sequence
of all prokaryotic promoters.\textsuperscript{{[}50, 51{]}} The
\selectlanguage{greek}σ\selectlanguage{english}\textsuperscript{70} factor of the host RNAP forms a specific
connection with the helix-turn-helix DNA binding motif of the -35
region. Thus, a mutation in this region could prevent the sigma factor
from binding to the promoter. The other three derivatives, B4.1, D2.4,
and E2.1, have a mutation in the \emph{lac} repressor LacI, indicating
the reduced productivity and increased growth rate. The reason for the
low productivity is the amino acid exchange in LacI, which was also
described to occur in Walker strain C43.\textsuperscript{{[}19,
22{]}}The mutations Q207H, Q146C, and D108E in \#B4.1, \#D2.1, and
\#E2.1, respectively, are located in the inducer binding pocket of LacI
and, thus, influence the binding affinity for IPTG and/or
allolactose\textsuperscript{{[}21{]}} (Figure S4). As a result, cells
have tuned themselves to the maximum tolerated productivity. This seems
to be a universal phenomenon for \emph{lac} -regulated gene expression.
Under completely different experimental conditions, a similar result was
achieved in Walker strain C43. Miroux and Walker produced mutants by
spreading cells on IPTG-containing agar. Although the cells were
cultured without process control, and despite the presence of the DE3
lysogen, the same type of variations developed in two independent
experiments. However, Miroux and Walker's experiments did not focus on
long-term stability, but on the productivity of toxic membrane proteins.
Based on our results, we conclude that T7 RNAP-dependent expression
systems acquire at least one mutation in the T7 RNAP gene and/or T7
promoter in long-term cultivation, regardless of the protein to be
produced.

\textbf{Table 2.} Mutations in Fab-producing derivatives of
B3\textless{}T7-Fab\textgreater{} and BQ\textless{}A1-Fab\textgreater{}
isolated after fractionation.\selectlanguage{english}
\begin{longtable}[]{@{}lllllll@{}}
\toprule
\textbf{Strain} & \textbf{Clone} & \textbf{Position} & \textbf{Mutation}
& \textbf{Region} & \textbf{Gene} & \textbf{Function}\tabularnewline
\midrule
\endhead
B3\textless{}T7-Fab\textgreater{} & C7 & 752679 & CGC-CTC (R756L) &
Coding & T7 RNAP gene & Enterobacteria phage T7 RNAP\tabularnewline
& D7 & 752266 & \selectlanguage{greek}Δ \selectlanguage{english}11 bp & Coding & T7 RNAP gene & Enterobacteria phage
T7 RNAP\tabularnewline
& F7 & 752331 & GGG-GAG (G640E) & Coding & T7 RNAP gene & Enterobacteria
phage T7 RNAP\tabularnewline
BQ\textless{}A1-Fab\textgreater{} & B4.1 & 336293 & CAG-CAT (Q207H) &
Coding & \emph{lacI} & Lac repressor\tabularnewline
& D2.4 & 336478 & GGC-TGC (G146C) & Coding & \emph{lacI} & Lac
repressor\tabularnewline
& E2.1 & 336590 & GAT-GAG (D108E) & Coding & \emph{lacI} & Lac
repressor\tabularnewline
& C3 & 3771324 & A - G & Intergenic & -96 from \emph{gfpmut3.1} &
\emph{pA1}\tabularnewline
\bottomrule
\end{longtable}

\textbf{3.3 Long-term stability of select derivatives in chemostat
cultivations}

The derivatives described above were generated under carbon-limited
fed-batch-like production conditions in the microbioreactor. This raised
the question about how select derivatives that can grow unhindered
despite recombinant protein production behave in long-term cultivations
and whether they can continue to maintain productivity over several
generations.

To address this question, we performed comparative lab-scale chemostat
cultivations. In the case of the easy-to-produce protein GFP, the best
mutant B3\textless{}T7-GFP\textgreater{} \#E7P6 S10, which had a
mutation in the pLac promoter, was compared to the non-mutated wild-type
strains BQ\textless{}A1-GFP\textgreater{} wt and
B3\textless{}T7-GFP\textgreater{} wt in long-term chemostat
cultivations.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image3/image3}

\end{center}
\end{figure}

\textbf{Figure 3.} Process characteristics of GFP-producing strains in
chemostat culture. (A-C) Courses of the specific content of GFP
(Y\textsubscript{P/X}) and biomass. (D-F) Single-cell expression
analysis. (A, D) B3\textless{}T7-GFP\textgreater{} wt, (B, E)
B3\textless{}T7-GFP\textgreater{} \#E7P6 S10, (C, F)
BQ\textless{}A1-GFP\textgreater{} wt.

For B3\textless{}T7-GFP\textgreater{} wt, induction with IPTG resulted
in very high specific content of 348 mg GFP/g CDM (Figure 3A), which was
clearly higher than the product titer from a fed-batch culture with the
same final biomass of 30~g/L (Figure 4A, Table 3). This productivity
could be maintained for 72 hours after induction. An extreme decrease
was observed in the biomass and product titer. This phenomenon was
demonstrated in previous studies in which we also observed a decrease in
the product titer 60 h after induction.\textsuperscript{{[}15{]}} After
approximately 120 hours, the biomass recovered and increased to the
intended 30 g/L CDM, but without product formation. The specific GFP
content remained at 0 mg/g until the end of the fermentation. As shown
in Figure 3D, non-producers had completely asserted themselves, and the
results of the microtiter experiments were reproduced. Similar behavior
was observed with the derivative \#E7P6 S10 (Figure 3B), with an extreme
decrease in the biomass to almost 0 g/L after 72 hours in chemostat
mode. After 120 hours, the cells recovered to the intended biomass of
30~g/L CDM. Unlike its wild-type ancestor, the specific GFP content did
not decrease to zero, but decreased steadily until the end of the
fermentation. The peak at a fluorescence intensity of
10\textsuperscript{2} in the FL1-A channel (Figure 3E) suggests that a
mixed population emerged in which non-producers have gradually overgrown
the producing cells, causing the production to decrease over the course
of the cultivation.

By determining the T7 RNAP gene sequence via PCR amplification of the
corresponding chromosomal region and Sanger sequencing, we identified
the insertion element insD-3 within the T7 RNAP gene of
B3\textless{}T7-GFP\textgreater{} wt and a mutation within the lacUV5
promoter in derivative \#E7P6 S10, which may have resulted in the
complete loss of GFPmut3.1 productivity

In contrast, no extreme decrease in biomass was observed for the host
RNAP-dependent expression system BQ\textless{}A1-GFP\textgreater{} wt,
indicating an extraordinarily high stability (Figure 3C). Over the
entire cultivation period of 312 hours and 45 generations, the specific
product content remained constant at approximately 100 mg GFP/g CDM
(Figure 3C). Flow cytometry confirmed the process stability. A
homogenous population was confirmed at each time point (Figure 3F). The
lower productivity of this expression system in combination with GFP
apparently represents such a low metabolic load that there was no
population collapse. Potential mutants did not have a growth advantage,
which means that the population of producing cells could persist for a
period of 45 generations.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image4/image4}

\end{center}
\end{figure}

\textbf{Figure 4.} Process characteristics of Fab-producing strains in
chemostat culture. Courses of the specific content of Fab
(Y\textsubscript{P/X}) and biomass. (A)
B3\textless{}T7-Fab\textgreater{} wt, (B)
BQ\textless{}A1-Fab\textgreater{}~wt, (C)
BQ\textless{}A1-Fab\textgreater{} \#D2.4, (D)
BQ\textless{}A1-Fab\textgreater{} \#E2.1.

Long-term chemostat cultivation with Fab were performed with two
BQ\textless{}A1-Fab\textgreater{} variants, \#D2.4 and \#E2.1 (Figure 4C
and D). The wild-type non-mutated strains
B3\textless{}T7-Fab\textgreater{} wt and
BQ\textless{}A1-Fab\textgreater{} wt served as reference (Figure 4A and
B).

Unlike GFP, the wild-type strain BQ\textless{}A1-Fab\textgreater{} wt
exhibited the same behavior as the strong T7 expression system
B3\textless{}T7-Fab\textgreater{} wt. In both wild-type strains, a
decrease in the biomass was already observed 6 hours after induction and
concomitant reduction of the product. The maximum amount of product was
4.9 mg/g and 2.5 mg/g for B3~\textless{}T7-Fab\textgreater{} wt and
BQ~\textless{}A1-Fab\textgreater{} wt, respectively. This corresponds to
the maximum measured product titers from fed-batch cultivations (Figure
4C and D, Table 3). Based on the results obtained for GFP, this behavior
was to be expected with the T7 expression system. Interestingly, the
phenomenon of the population collapse was observed using the
BQ\textless{}A1\textgreater{} expression system. As mentioned above, the
degree of metabolic load depends not only on the expression system, but
also on the specific recombinant protein to be produced. In the case of
Fab, the expression of even the weaker promoter in
BQ\textless{}A1-Fab\textgreater{} wt was too strong. Sanger sequencing
of the A1 promoter again revealed an A -\textgreater{} G substitution
within the -35 region. In contrast, no notable decrease in biomass was
observed in the two A1 derivatives. Derivative \#D2.4 was able to
consistently produce approximately 1.0 mg Fab/g CDM over a total of 120
hours, which corresponds to 17.3 generations. Derivative \#E2.1 produced
even more at 1.6 mg/g. Nevertheless, long-term stability was evidently
accompanied by a reduction in productivity. Compared to the conventional
fed-batch processing, in which BQ\textless{}A1-Fab\textgreater{} wt
produced 2.7 mg/g (Figure 4D), the product titers of the corresponding
derivatives were clearly lower, approximately half in the case of the
best producing mutant \#E2.1.

\textbf{Table 3.} Comparison of productivity between fed-batch and
chemostat cultivations (final biomass 30 g/L CDM).\selectlanguage{english}
\begin{longtable}[]{@{}lllll@{}}
\toprule
\textbf{Strain} & \textbf{Cultivation mode} & \textbf{Max. volumetric
yield {[}g/L{]}\textsuperscript{a}} & \textbf{Process
time~\textsuperscript{b} {[}h{]}} & \textbf{STY
{[}mg/L/h{]}~\textsuperscript{c}}\tabularnewline
\midrule
\endhead
B3\textless{}T7-GFP\textgreater{} wt & Fed-batch & 8.3 \selectlanguage{ngerman}± 0.6 & 29 &
286.2\tabularnewline
B3\textless{}T7-GFP\textgreater{} wt & Chemostat & 10.9 &
60~\textsuperscript{d} & 847.1\tabularnewline
BQ\textless{}A1-GFP\textgreater{} wt & Fed-batch & 1.8 \selectlanguage{ngerman}± 0.2 & 29 &
62.8\tabularnewline
BQ\textless{}A1-GFP\textgreater{} wt & Chemostat & 2.6 & 324 &
284.1\tabularnewline
B3\textless{}T7-GFP\textgreater{} \#E7P6 S10 & Chemostat & 10.2 &
60~\textsuperscript{d} & 642.2\tabularnewline
B3\textless{}T7-Fab\textgreater{} wt & Fed-batch & 0.052 \selectlanguage{ngerman}± 0.006 & 29 &
1.8\tabularnewline
B3\textless{}T7-Fab\textgreater{} wt & Chemostat & 0.164 & 18 &
3.4\tabularnewline
BQ\textless{}A1-Fab\textgreater{} wt & Fed-batch & 0.087 \selectlanguage{ngerman}± 0.008 & 29 &
3.0\tabularnewline
BQ\textless{}A1-Fab\textgreater{} wt & Chemostat & 0.085 & 18 &
1.8\tabularnewline
BQ\textless{}A1-Fab\textgreater{} \#E2.1 & Chemostat & 0.035 & 132 &
4.3\tabularnewline
BQ\textless{}A1-Fab\textgreater{} \#D2.4 & Chemostat & 0.019 & 132 &
2.3\tabularnewline
\bottomrule
\end{longtable}

\textsuperscript{a} For fed-batch cultivations (Figure S5), the mean \selectlanguage{ngerman}±
standard deviation for individual determinations is given (n=3).

\textsuperscript{b} Process time is the sum of reactor setup and the
actual fermentation

\textsuperscript{c} Space time yield (STY) calculated according to
Equation 1.

\textsuperscript{d} Chemostat cultivations in which the cultivation
period was limited to the phase before population collapse.

If long-term stability is only possible through reduced productivity,
the question of process efficiency arises. In order to compare fed-batch
and continuous production, we used the space-time-yield (STY; Equation
1) as the evaluation criterion.\selectlanguage{english}
\begin{longtable}[]{@{}ll@{}}
\toprule
\(STY=\frac{\frac{\text{Total\ protein\ }\left[\text{mg}\right]}{\text{Reactor\ volume\ }\left[L\right]}}{\text{Process\ time\ }\left[h\right]}\) & (1)\tabularnewline
\bottomrule
\end{longtable}

For calculation of the process time for both process types, we added a
down time of 12 hours for media preparation, reactor setup, CIP, and SIP
to the cultivation time. In chemostat cultivations with population
collapse, we only considered the cultivation time before the decrease in
cell density, which was the case with B3 \textless{}T7-GFP\textgreater{}
after 48 hours.

As can be concluded from the productivity comparisons (Table 3), the
fed-batch process with the T7 expression system and the model protein
GFP were inferior to the chemostat process in terms of volumetric yield
and STY. The reason for this can be attributed to the duration of the
induction. In fed-batch cultivation, the production phase takes 19
hours. Chemostat cultivation can be extended to 48 hours because the
volume in the reactor is kept constant.

As already mentioned, a longer chemostat process would not be possible
under these circumstances because the process ultimately becomes
unstable. However, a short-term chemostat cultivation of 48 hours could
be interesting for easy-to-produce proteins. With 874 mg/L/h, the STY of
B3\textless{}T7-GFP\textgreater{}wt in the chemostat tripled compared to
the fed-batch cultivation. In the case of the host RNAP-dependent A1
expression system, the STY of 62.8 mg/L/h in fed-batch cultivation was
relatively low compared to the STY of 286 mg/L/h of the conventional T7
expression system. In addition to the short production time of 19 hours,
the A1 expression system was inherently weaker. However, the low titer
can be compensated by continuous production. Due to the very long stable
production time of 312 hours, the STY of the chemostat cultivation more
than quadrupled compared to the fed-batch cultivation. Nevertheless, for
the production of GFP, the conventional T7 expression system is still
superior overall, in both fed-batch and chemostat mode, especially when
considering short-term chemostat cultivation of the T7 wild-type strain
(Table 3).

The situation is different with Fab. The extraordinary strength of the
T7 expression system is a disadvantage for Fab production. In neither
fed-batch nor chemostat mode, the T7 expression system was able to
outperform the host RNAP-dependent A1 expression systems. Due to the
mutation in \emph{lacI} , the volumetric yield of
BQ\textless{}A1-Fab\textgreater{}wt was reduced from 0.087 g/L to 0.035
g/L, as seen with derivative \#E2.1. However, long-term cultivation
without population collapse was possible. Due to the long production
time of 120 h, the STY of the BQ\textless{}A1-Fab\textgreater{}wt was
increased from 3.00 g/L/h in fed-batch mode to 4.33 g/L/h in chemostat
mode for derivative \#E2.1 when comparing the two strains to one
another.

These results indicate that, in continuous \emph{E. coli} bioprocesses,
the choice of expression system depends on the recombinant protein to be
produced. In the case of an easy-to-produce protein, such as GFP, the
conventional T7 expression system is still the expression system of
choice. The duration of the chemostat cultivation is limited by the
inevitable population collapse, but this can be compensated by the
extraordinary strength of the T7 system.

However, the moderately strong host RNAP-dependent A1 system is more
suitable for the production of a challenging protein. As a result of the
large metabolic load that is triggered, a reduction in the expression
rate is advantageous. Thus, the low titer can not only be compensated,
but even increased, based on the STY. As already mentioned, the time in
which the actual fermentation occurs and recombinant protein is produced
in fed-batch cultivation is relatively low. In addition, the cost of CIP
and SIP must be included. Although these process steps must also be
carried out in a long-term chemostat cultivation, the phase in which the
actual production occurs is inevitably prolonged, which means that the
CIP and SIP frequency can be reduced.

Nevertheless, only mutants that have adapted to long-term stability
under production conditions can be cultivated in long-term chemostat
mode. In wild-type strains, the population collapse and adaptive rescue
during long-term chemostat cultivation does not allow long-term
production. This phenomenon has been observed frequently in
other\emph{E. coli} chemostat cultivations and prevents the direct use
of non-adapted strains for this purpose.\textsuperscript{{[}5, 52,
53{]}}

\textbf{4 Concluding remarks}

Identification of genetically stable \emph{E. coli} mutants using
high-throughput serial fed-batch microtiter cultivations was
successfully implemented. Here, we reported a selection-based approach
based on the growth rate under production
conditions.\textsuperscript{{[}23, 24{]}} As expected, the genetic
stability of a T7 expression system is not sufficient to maintain
constant product formation levels during long-term cultivation under
production conditions. Mutations leading to non-producers are located in
the T7 RNAP gene and/or the T7 promoter. The host RNAP-based A1
expression system, which has moderate expression strength, remained
stable in the production of the easy-to-produce protein GFP during
long-term cultivation. For the production of challenging proteins (e.g.,
Fabs), which trigger more severe metabolic load on cell metabolism,
mutations in \emph{lacI} of BQ\textless{}A1-Fab\textgreater{}
derivatives reduce the expression levels but have positive effects on
long-term stability. We could not find any mutations with positive
effects on protein expression, such as mutations in folding helpers, the
Sec translocon, or proteases that could serve as modification targets in
a rational approach. This finding was not unexpected, as no selection
pressure was applied to production of the recombinant protein.

In addition to obvious mutations in the T7 RNAP gene and/or T7 promoter
that led to reduced burden and higher growth rates, we were able to find
mutations in the metabolism and transport mechanism of various sugars.
These mutations led most to decreased inducer uptake and, thus, reduced
induction. However, three mutants were found with unrestricted growth
and stable Fab expression at a reduced, but obviously physiologically
acceptable, level. For continuous recombinant protein production
in\emph{E. coli} , cells have to adapt to their maximum tolerable level
of recombinant protein production. Therefore, the mutations found here
do not generally apply to all challenging or easy-to-produce recombinant
proteins, but can vary from protein to protein.

Regarding industrial regulations, the US FDA regulatory body encourages
the biological industries to use continuous manufacturing approaches for
the production of new products. This aspect applies to the entire
bioprocess, including downstream processing. Thus far, continuous
downstream processing has been more relevant to mammalian perfusion
processes.\textsuperscript{{[}12{]}} However, with the production
strains described in this study, \emph{E. coli} processes could also be
relevant.

In conclusion, we have shown that, through a directed evolution
approach, a high- throughput screening process comes very close to
industrial production processes, and we were able to find derivatives
that have a positive effect on long-term stability. We postulate that
long-term stability studies with \emph{E. coli} can only be carried out
with genome-integrated expression systems. As plasmid loss is no longer
a problem in this regard, we were able to characterize how metabolic
load triggered by recombinant protein production influences the
characteristics of mutations in \emph{E. coli} . Thus, adaptive
evolution in microtiter cultivations could be an efficient strain
development method in addition to a rational approach. Although the
specific product titers are reduced, they can be compensated by
continuous production.

\textbf{Acknowledgement}

We thank Philipp Peters for data analysis at an early stage of the
project and Nancy Stralis-Pavese for preparation of the sequencing
libraries. Some calculations were performed at the Vienna Scientific
Cluster (VSC). Illumina sequencing was performed at the Vienna BioCenter
Core Facilities (VBCF) Next Generation Sequencing Facility.

The financial support by the Austrian Federal Ministry for Digital and
Economic Affairs, the National Foundation for Research, Technology and
Development and the Christian Doppler Research Association is gratefully
acknowledged.

Furthermore, we gratefully acknowledge the financial support received
from Boehringer Ingelheim RCV GmbH \& Co KG. The funding agencies had no
influence on the conduct of this research. We thank the colleagues of our
company partner BI RCV for the scientific input and their continuous
scientific support.

\textbf{Author contributions:}

AS, MCP, RG, JCD, HH, and GS designed the research; AS, CK, and MS
performed the research; AS, MCP, RG, MW, JCD, MS, HH, and GS analyzed
the data; AS and GS wrote the paper, with contributions from MS, JCD,
MCP, and MW. All authors approved the final version of the manuscript.

\textbf{Conflict of interest}

The authors declare no commercial or financial conflict of interest.

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