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\begin{document}

\title{Uncoupling growth and succinic acid production in an industrial
\emph{Saccharomyces cerevisiae} strain}



\author[1]{Yaya Liu}%
\author[1]{Osman Esen}%
\author[1]{Jack T. Pronk}%
\author[1]{Walter Martin van Gulik}%
\affil[1]{TU Delft}%


\vspace{-1em}



  
  \date{May 18, 2024}


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\selectlanguage{english}
\begin{abstract}
This study explores the relation between biomass-specific succinic acid
(SA) production rate and specific growth rate of an engineered
industrial strain of \emph{Saccharomyces cerevisiae}, with the aim to
investigate the extent to which growth and product formation can be
uncoupled. Ammonium-limited aerobic chemostat and retentostat cultures
were grown at different specific growth rates under industrially
relevant conditions, i.e., at a culture pH of 3 and with sparging of a
1:1 CO2-air mixture. Biomass-specific SA production rates decreased
asymptotically with decreasing growth rate. At near-zero growth rates,
the engineered strain maintained a stable biomass-specific SA production
rate for over 500 h, with a SA yield on glucose of 0.61
mol.mol\textsuperscript{-1}. These results demonstrate that uncoupling
of growth and SA production could indeed be achieved. A linear relation
between biomass-specific SA production rate and glucose consumption rate
indicated a coupling of SA production rate and the flux through primary
metabolism. The low culture pH resulted in an increased death rate,
which was lowest at near-zero growth rates. Nevertheless, a significant
amount of non-viable biomass accumulated in the retentostat cultures,
thus underlining the importance of improving low-pH tolerance in further
strain development for industrial SA production with \emph{S.
cerevisiae.}%
\end{abstract}%



\sloppy


\par\null

\subsection*{Introduction}

{\label{introduction}}

Succinic acid (SA), a C4 dicarboxylic acid and intermediate of the
tricarboxylic acid (TCA) cycle (Tretter, Patocs, \& Chinopoulos, 2016),
is used as precursor for manufacturing a wide range of products in the
pharmaceutical, agricultural, and food industries, including detergents,
fungicides, herbicides, biodegradable polymers, flavors and food
additives (Zeikus, Jain, \& Elankovan, 1999). Based on predicted
environmental benefits, 2004 and 2010 reports of the U.S. Department of
Energy (DOE) mentioned SA as one of the five most promising bio-based
chemicals (Bozell \& Petersen, 2010; Werpy \& Petersen, 2004).The SA
market size is expected to reach 710,000 tons by 2020, with a predicted
turn-over of USD 1.1--1.3 billion in 2022 (Dessie et al., 2018; Luthfi
et al., 2017). In order to supply this market, bio-based production of
SA will become increasingly important as it can provide a sustainable
alternative to petrochemical production (Pinazo, Domine, Parvulescu, \&
Petru, 2015).

In recent years, production of SA in industrial fermentation processes
has been realized with naturally SA-producing microorganisms as well as
with engineered strains (Beauprez, De Mey, \& Soetaert, 2010; Chen \&
Nielsen, 2013). BASF/Corbion-Purac achieved a yield of 0.75 mol of SA
per mol of glucose from the natural producer \emph{Basfia
succiniproducens} , which was isolated from bovine rumen (Kuhnert,
Scholten, Haefner, Mayor, \& Frey, 2010). Myriant applied an
engineered\emph{E.coli} strain for large-scale SA production (Ahn, Jang,
\& Lee, 2016). However, SA-producing bacteria may be affected by
bacteriophage infection and generally require a neutral culture pH. The
latter poses a requirement for alkali titration during fermentation and
a subsequent acidification during downstream processing, resulting in
massive waste production in the form of gypsum (Abbott, Zelle, Pronk, \&
van Maris, 2009). Fermentation at a low pH provides a way to reduce
waste production and, thereby, improve process economics and
sustainability.

Due to their low pH tolerance and insensitivity to phage infection,
engineered yeasts have been intensively studied as microbial SA
production platforms. For instance, Bioamber/Mitsui has replaced its SA
producing \emph{E.coli} strain with the yeast \emph{Candida
krusei}(Jansen \& van Gulik, 2014). Since 2012\emph{,} DSM/Roquette has
applied an engineered \emph{S. cerevisiae} strain for industrial scale
SA production (Jansen, Heijnen, \& Verwaal, 2013). To reach a high SA
yield, the latter strain was genetically modified by overexpression of
the reductive branch of the TCA cycle in the cytosol, with further
genetic modification focused on the glyoxylate cycle and SA export
across the plasma membrane. Because SA production via the reductive part
of the TCA cycle involves a net consumption of CO\textsubscript{2},
increasing dissolved CO\textsubscript{2} concentrations should increase
the driving force for SA biosynthesis via this pathway. It has indeed
been reported that enrichment of the aeration gas with
CO\textsubscript{2} significantly increased the rate of SA production in
engineered strains of \emph{S. cerevisiae} (Jamalzadeh, 2013; Shah,
2016; Zelle, de Hulster, Kloezen, Pronk, \& van Maris, 2010). Although
the Gibbs free energy changing of the synthesis of SA from glucose is
negative, the costs of active SA export (2 to 3 mol ATP per mol SA
exported) (Taymaz-Nikerel et al., 2013) require a net input of ATP.
Oxygen therefore needs to be supplied during yeast-based SA production
to enable ATP production via respiration (Shah, 2016).

In a study on product recycling across yeast membranes at high SA
titers, which was based on a scaled-down industrial SA fed-batch
fermentation process at low pH (Wahl, Bernal Martinez, Zhao, van Gulik,
\& Jansen, 2017), significant \textsuperscript{13}C labeling of the TCA
cycle intermediates fumarate, isocitrate and \selectlanguage{greek}α-\selectlanguage{english}ketoglutarate was
observed within 100 s after extracellular addition
of\textsuperscript{13}C labelled SA. This observation indicated that SA
rapidly exchanges over the plasma membrane. In this scaled-down
fed-batch process, SA production rates declined with decreasing specific
growth rate. This observation was attributed to increased product
degradation as well as to increased non-growth associated energy
requirements at high SA titers, which left less substrate available for
energy-dependent product formation.

From an industrial point of view, fermentative production of SA in the
absence of cell growth would be ideal, as it would minimize formation of
biomass as a byproduct and maximize the yield of product on substrate.
Such an uncoupling of growth and product formation requires a producing
strain which can maintain a high productivity at near-zero specific
growth rates. Due to their dynamic nature, fed-batch cultures are not
the best option to study relations between specific growth rate and
strain performance (Hewitt \& Nienow, 2007). In contrast, chemostat
cultivation allows studies on microbial physiology at a constant
specific growth rate, under well-defined, stable process conditions.
However, chemostat cultivation in laboratory bioreactors is impractical
at specific growth rates below 0.025 h\textsuperscript{-1}, due to the
long time periods needed to reach steady state. As an alternative to
chemostat cultivation, retentostat cultures have proven to be excellent
tools to study growth of \emph{S. cerevisiae} and other microorganisms
at low to near-zero growth rates under various nutrient limitations
(Boender, de Hulster, van Maris, Daran-Lapujade, \& Pronk, 2009; Ercan
et al., 2015; Hakkaart et al., 2020; Liu, El Masoudi, Pronk, \& van
Gulik, 2019; Vos et al., 2016).

The goal of the present study is to investigate whether uncoupling of
growth and product formation can be accomplished in a heavily
engineered, SA high-producing industrial \emph{S. cerevisiae} strain.
The strain was grown under industrially relevant conditions, i.e. at an
elevated CO\textsubscript{2} level to increase the driving force towards
SA biosynthesis via the reductive pathway and a culture pH of 3.0 to
facilitate downstream processing (Hakkaart et al., 2020).
Ammonium-limited cultures were used, in which glucose was present in
excess, to avoid competition for glucose between growth and product
formation. Ammonium-limited chemostat and retentostat cultures were used
to the physiology of the industrial strain over a range of low to
near-zero specific growth rates. This approach enabled a quantitative
assessment of the degree to which biomass-specific production SA depends
and specific growth rate are coupled and to identify goals for further
strain engineering to improve uncoupling of growth and product formation
in industrial SA production.

\subsection*{Materials and methodsYeast strain and growth
media}

{\label{697253}}

The engineered SA-overproducing industrial \emph{S. cerevisiae} strain
(SUC632) was kindly provided by Royal DSM B.V (Delft, the Netherlands).
Properties and performance of this strain have been described previously
(Jansen et al., 2013; Wahl et al., 2017). Working stocks were stored at
-80 \selectlanguage{ngerman}\textbf{°} C in 1 mL aliquots in YPD medium (10 g/L~Bacto yeast
extract, 20 g/L~Bacto peptone, 20 g/L~glucose) containing 30 \% (v/v)
glycerol. The pre-culture medium contained, per liter of demineralized
water: 20 g galactose, 2.3 g urea, 3.0 g
KH\textsubscript{2}PO\textsubscript{4} , 0.5 g
MgSO\textsubscript{4}.7H\textsubscript{2}O, 1 ml trace-element solution
and 1 ml vitamin solution (Verduyn, Postma, Scheffers, \& Van Dijken,
1992). After filter sterilization (Millex® Syringe Filters, 0.22 \selectlanguage{greek}µ\selectlanguage{english}m,
Merck Millipore, Massachusetts,~USA), 1 ml of sterilized chalk solution
(0.1 g/g CaCO\textsubscript{3}) was added per 100 ml pre-culture medium.
The medium for batch cultivation contained, per liter of demineralized
water: 30 g galactose, 1.0 g
(NH\textsubscript{4})\textsubscript{2}SO\textsubscript{4}, 5.3 g
K\textsubscript{2}SO\textsubscript{4}, 3.0 g
KH\textsubscript{2}PO\textsubscript{4} , 0.5 g
MgSO\textsubscript{4}.7H\textsubscript{2}O, 1 ml trace element solution,
1 ml vitamin solution, 1 ml of a solution containing 3 g/L
FeSO\textsubscript{4}.7H\textsubscript{2}O and 15 g/L EDTA, 1 ml of
biotin solution (1 g/L), and 0.2 g Pluronic 6100 PE antifoaming agent
(BASF, Ludwigshafen, Germany). The composition of the chemostat feed
medium was the same as the medium used for the batch phase, except for
the carbon source, which was 30 g/L glucose instead of 30 g/L galactose.
For retentostat cultivation concentrations of glucose and
(NH\textsubscript{4})\textsubscript{2}SO\textsubscript{4} were decreased
to 18 g/L and 0.1 g/L, respectively.

\subsubsection*{Pre-cultures and aerobic bioreactor
cultures}

{\label{pre-cultures-and-aerobic-bioreactor-cultures}}

Pre-cultures were initiated by inoculating 500 mL Erlenmeyer flasks
containing 200 mL of pre-culture medium with 2 mL of stock culture and
incubated for 24 h in an orbital shaker at 30\selectlanguage{ngerman}°C and at a rotation speed
of 200 rpm (B Braun Certomat BS-1, Sartorius, Melsungen, Germany).

Batch, chemostat and retentostat cultivations were carried out in 7 L
bioreactors (Applikon, Delft, The Netherlands) as described previously
(Liu et al., 2019).

During batch cultivation, bioreactors were sparged with compressed air
(0.04 \% CO\textsubscript{2}). During chemostat and retentostat
cultivation, the reactors were sparged with a 1:1 mixture of air and
CO\textsubscript{2} (\textgreater{} 99.7\% purity, Linde Gas Benelux,
Schiedam, The Netherlands) by using two mass-flow controllers. The pH
was maintained at 3.00 ± 0.05 by automatic titration with 2 M KOH or 2 M
H\textsubscript{2}SO\textsubscript{4}. The dissolved-oxygen tension was
measured using an autoclavable Clark sensor (Mettler-Toledo GmbH,
Greinfensee, Switzerland) but was not controlled. Exhaust gas from
bioreactors was dried by first passing through a condenser at 4°C and
then through a Perma Pure Dryer (Inacom Instruments, Overberg, The
Netherlands). CO\textsubscript{2} and O\textsubscript{2} concentrations
in the dried gas were measured with a Rosemount NGA 2000 gas analyzer
(Minnesota, USA).

Independent duplicate chemostat cultures were carried out at dilution
rates of 0.07, 0.058, 0.048, 0.035 and 0.025 h\textsuperscript{-1}.
Steady state was assumed to be achieved when off-gas
CO\textsubscript{2}and O\textsubscript{2} concentrations, biomass dry
weight and cell counts changed by less than 3 \% difference over 2
consecutive volume changes. After these criteria had been met, samples
were withdrawn after 5, 6 and 7 subsequent volume changes for
measurement of biomass dry weight, cell number, concentrations of
residual substrates and (by)products and culture viability.

Retentostat cultures were started with a batch cultivation phase,
followed by chemostat cultivation at a dilution rate of 0.025
h\textsuperscript{-1}. After reaching steady state, chemostat cultures
were switched to retentostat mode by switching off the chemostat
broth-removal system and activating cell-free culture removal through
four filtration probes (Applikon, Delft, The Netherlands) mounted inside
the reactor (Liu et al., 2019). The cell-free effluent was pumped in a
sterile effluent vessel using a peristaltic pump (Masterflex,
Barrington, USA) controlled by the vessel weight, such that the broth
weight was kept at 5.00 ± 0.05 kg. During retentostat cultivation,
dilution rate was maintained at 0.025 h\textsuperscript{-1} . A gradual
transition from chemostat to retentostat medium was accomplished by
using two feed pumps (Liu et al., 2019). Duplicate retentostats were
carried out and samples were withdrawn every 48 h until the cultures
were terminated.

\subsubsection*{Determination of biomass dry weight, cell counts and
culture
viability}

{\label{determination-of-biomass-dry-weight-cell-counts-and-culture-viability}}

Biomass dry weight was quantified gravimetrically. Total cell counts
were measured by a Z2 Coulter counter (50 \selectlanguage{greek}μ\selectlanguage{english}m aperture, Beckman,
Fullerton, CA). Cell viability of culture samples was determined with
a~FungaLight Yeast CFDA, AM/Propidium Iodide Vitality Kit and flow
cytometry. Detailed descriptions of the dry weight, cell counts and
viability measurements were described previously (Liu et al., 2019).

\subsubsection*{Rapid sampling for metabolite
quantification}

{\label{rapid-sampling-for-metabolite-quantification}}

Cell-free effluent samples for quantification of extracellular
metabolites were obtained from a sample port connected to the
retentostat filter assembly. Broth samples for intracellular metabolite
measurements were withdrawn by a rapid sampling device connected to the
bioreactor as described previously (Liu et al., 2019).

\subsubsection*{Quantification of substrates, products and intracellular
metabolites}

{\label{quantification-of-substrates-products-and-intracellular-metabolites}}

Ammonium concentrations were analysed with a Gally Discrete Analyzer
(ThermoFisher Scientific, Massachusetts, United States) with a detection
limit of 0.02 mg NH4\textsuperscript{+}/L. Concentrations of glucose,
succinic acid and by-products (malate, ethanol, glycerol, acetate and
lactate) were quantified by HPLC, using a Bio-Rad HPX-87H 300 column
(7.8 mm) as described previously (Liu et al., 2019). Intracellular
metabolite concentrations were quantified using isotope dilution mass
spectrometry (LC-IDMS/MS and GC-IDMS) with
U-\textsuperscript{13}C-labeled yeast cell extract as internal standard
(Wu et al., 2005). Detailed descriptions of the mass spectrometry based
quantification protocols were published previously (Cipollina et al.,
2009; Seifar et al., 2009; van Dam et al., 2002). The adenylate energy
charge (AEC) was calculated as described (Liu et al., 2019).

\subsubsection*{Biomass-specific rate
calculations}

{\label{biomass-specific-rate-calculations}}

Calculation of specific growth rates (\selectlanguage{greek}µ, \selectlanguage{english}h\textsuperscript{-1}) and
specific death rates (k\textsubscript{d}, h\textsuperscript{-1}) were
performed as described previously (Liu et al., 2019). Biomass-specific
glucose and ammonium consumption rates and biomass-specific production
rates of succinic acid, malate, ethanol, glycerol, acetate and lactate
were calculated from the primary measurements of substrate and product
concentrations and flow rates in gas and liquid phases using the
corresponding material balances. Data reconciliation was performed as
described by (Verheijen, 2000).

\subsection*{Results}

{\label{285966}}

Growth and viability of S. cerevisiae SUC632 in ammonium-limited
chemostat and retentostat cultures at low pH and high
CO\textsubscript{2}levels

To investigate whether the specific SA production rate of the
engineered\emph{S. cerevisiae} strain SUC632 was related to its specific
growth rate, the strain was cultivated under industrially relevant
conditions (50 \% CO\textsubscript{2}, pH 3.0) in ammonium-limited
cultures. Specific growth rates between 0.034 and 0.085
h\textsuperscript{-1} were studied in chemostat cultures, while
retentostat cultures were applied to obtained quantitative data on
strain performance at specific growth rates from 0.034
h\textsuperscript{-1} to near zero. All these continuous cultures were
grown under ammonium limitation. In all retentostat and steady state
chemostat cultures, concentrations of residual ammonium were below the
detection limit, and residual glucose concentrations were higher than
the saturation constant of the high-affinity glucose transporters in
\emph{S. cerevisiae}(k\textsubscript{m}, ca. 1 mM) (Boles \& Hollenberg,
1997; Reifenberger, Boles, \& Ciriacy, 1997), indicating that ammonium
was indeed the growth limiting nutrient (Supporting Information Table
S1).

In all chemostat cultures, viability was between 80 and 90 \% (Figure
1A). As only the viable cell fraction contributes to substrate
consumption, growth and (by)product formation, specific conversion rates
were expressed per g of viable cells. The steady-state total and viable
biomass concentrations slightly increased with decreasing specific
growth rate to become stable below a specific growth rate of 0.042
h\textsuperscript{-1} (Figure 1C). A similar profile was observed for
total cell counts, while the average dry mass per cell decreased with
decreasing specific growth rate (Figure 1E).

As, during retentostat cultivation, the specific growth rate
progressively decreased from 0.025 h\textsuperscript{-1} to near zero,
culture viability declined from 80 \% to a stable value of around 25 \%
between 300 and 450 h of cultivation, with a slight further decrease
thereafter (Figure 1B). Total biomass dry weight and cell counts
increased during retentostat cultivation according to similar patterns
Figure 1D, F). The viable biomass concentration levelled off at 2 g/L
after 300 h (Figure 1D). The dry mass per cell was stable during
retentostat cultivation (Figure 1F) and identical to the value observed
in chemostat cultures grown at 0.034 h\textsuperscript{-1}.

The highest specific death rate (k\textsubscript{d} = 0.015
h\textsuperscript{-1}) was observed at the highest specific growth rate
of 0.085 h\textsuperscript{-1} applied during chemostat cultivation. In
the other, slower growing chemostat cultures, k\textsubscript{d} was
approximately 0.01 h\textsuperscript{-1} (Figure 1G). From the start of
the retentostat cultivations, k\textsubscript{d} decreased from 0.01
h\textsuperscript{-1} to approximately 0.005 h\textsuperscript{-1},
meanwhile the corresponding specific growth rates (\selectlanguage{greek}μ\selectlanguage{english}) also rapidly
dropped and approximately stabilized at values below 0.005
h\textsuperscript{-1} (Figure 1H).

\subsubsection*{Uncoupling of growth and succinic acid
production}

{\label{uncoupling-of-growth-and-succinic-acid-production}}

In nitrogen-limited retentostat cultures, the biomass-specific SA
production rate decreased asymptotically with decreasing specific growth
rate and stabilized at a value of 0.75 mmol/(g viable biomass)/h for
specific growth rates between 0.008 and 0.014
h\textsuperscript{-1}(Figure 2). This result showed that the engineered
strain was still capable of producing SA at near-zero specific growth
rates. Apart from SA, malate, ethanol, glycerol and small amounts of
acetate and lactate were formed as byproducts. The biomass-specific
production rates of these by-products were combined in a
``q\textsubscript{byp}'' term and expressed as mCmol per gram of viable
biomass per h. The specific rate of byproduct formation decreased
asymptotically with decreasing specific growth rate to relatively low
values (1.2 mCmol/(g viable biomass)/h, Figure 2C). Individual
production rates for each by-product are provided in Supporting
Information Table S2.

The fraction of consumed glucose distributed to biomass, SA and
byproducts formation were compared between those different specific
growth rates cultures (Figure 3). Specifically, the fraction of the
consumed glucose converted to SA was highest in the condition that
growth was virtually absent (Figure 3). Besides, during retentostat
cultivation, the yield of SA on glucose progressively decreased from
almost 0.9 mol/mol until, after 400 h, it reached a stable value of
approximately 0.6 mol/mol (Figure 4).

\subsubsection*{Cellular energy status during near-zero growth
cultivation}

{\label{cellular-energy-status-during-near-zero-growth-cultivation}}

To investigate the energy status of the SA-producing strain at near-zero
growth rates, under the severe nitrogen limitation in low-pH retentostat
cultures, intracellular levels of adenine nucleotides (ATP, ADP and
AMP), and the adenylate energy charge and ATP/ADP ratios were analysed
(Figure 5). During retentostat cultivation, intracellular levels of ATP
and ADP slowly decreased (Figure 5B and C) and stabilized after about
350 h. Retentostat cultures showed a stable intracellular AMP level that
approximately two-fold higher than observed in the preceding chemostat
culture (Figure 5A). Despite these differences in the concentrations of
individual adenine nucleotides, the energy charge did not change upon
the switch from chemostat to retentostat cultivation and remained at a
stable value of approximately 0.8 throughout the retentostat experiments
(Figure 5D). The ATP/ADP ratio slowly increased from a value of 1.5 to
2.0 at the end of the retentostat cultivation (Figure 5E).

\subsubsection*{Levels of the TCA cycle metabolites during retentostat
cultivation}

{\label{levels-of-the-tca-cycle-metabolites-during-retentostat-cultivation}}

The target product in this study, SA, as well as the most important
by-product, malate, are both intermediates of the TCA cycle. To assess
how SA production correlated with intracellular levels of other
TCA-cycle intermediates, intracellular metabolite analyses were
performed on the retentostat cultures (Figure 6A-F). Levels of all
quantified TCA-cycle intermediates increased during retentostat
cultivation of the engineered strain, but followed distinct dynamic
patterns. Intracellular levels of citrate, isocitrate and succinate
increased linearly with time during retentostat cultivation, levels of
\selectlanguage{greek}α-\selectlanguage{english}ketoglutarate and fumarate decreased until 150 h and increased
thereafter, while the malate level increased until 300 h and then
decreased. Intracellular levels of citrate, succinate and malate were
two orders of magnitude higher than those of other TCA-cycle
intermediates. When compared to a wild-type strain grown at similar
glucose consumption rates in aerobic, glucose-limited chemostat cultures
(Canelas, Ras, ten Pierick, van Gulik, \& Heijnen, 2011) levels of
citrate, succinate and malate were 5, 90 and 25 fold higher,
respectively, towards the end of the nitrogen-limited retentostat
cultures. Remarkably, the mass action ratio of the fumarase reaction
(Figure 6G) was far above the apparent \emph{in vivo} equilibrium
constant of 5.2 (Canelas et al., 2011). This observation suggests that
the overall flux was from malate to fumarate, with a low fumarate level
resulting from the action of the heterologous, cytosolically expressed
NADH-dependent fumarate reductase (FRDg) from \emph{Trypanosoma
brucei}(Jansen et al., 2013). Succinate formation via this reaction is
highly exergonic, thus providing a plausible explanation for the high
intracellular succinate and low fumarate levels. The expression of
isocitrate lyase (KIICL1) from \emph{Kluyveromyces lactis} (Jansen et
al., 2013) may be responsible for the high intracellular citrate levels
as this enzyme, together with aconitase, allows for conversion of
conversion of succinate via isocitrate to citrate and vice versa. The
observation that the mass action ratio for the aconitase reaction was
well below the estimated \emph{in vivo} K\textsubscript{eq}, (Figure
6H), indicate that net flux through this enzyme was in the direction of
isocitrate.

\subsection*{Discussion}

{\label{discussion}}

In this study, we explored the feasibility of applying cultivation at
near-zero growth rate for high-yield production of chemicals from
renewable feedstocks. As a model system we used an engineered industrial
strain of \emph{S. cerevisiae} , capable of producing SA from glucose.
To simulate industrial process conditions, growth studies were performed
at low pH and at an elevated CO\textsubscript{2} concentration.

Fermentation processes at near-zero growth rate theoretically allow for
highly efficient conversion of substrate to product, as no carbon- and
energy source is required for biomass formation. However, long-term
cultivation of \emph{S. cerevisiae} at near-zero growth rate in
retentostat cultures has been reported to result in accumulation of
non-viable, non-producing cells (Boender et al., 2009; Vos et al.,
2016). A previous study with the non-producing laboratory reference
strain CEN.PK113-7D (Hakkaart et al., 2020) showed that, at pH 3, the
specific death rates (k\textsubscript{d}) in glucose- and
ammonium-limited retentostat cultures (respectively 0.0039 \selectlanguage{ngerman}± 0.0005 and
0.0030 ± 0.0004 h\textsuperscript{-1}) were approximately eight-fold
higher than in corresponding cultures grown at pH 5. However,
cultivation of the reference strain at near-zero growth rate as such did
not cause an increased k\textsubscript{d}, since the death rates in
glucose- and ammonium-limited retentostat cultures performed at pH 3
were two-fold and tenfold lower, respectively, than in corresponding
chemostat cultures grown at a specific growth rate of 0.025
h\textsuperscript{-1} (Hakkaart et al., 2020).

In the present study, ammonium-limited steady-state chemostat
cultivation at a specific growth rate of 0.025 h\textsuperscript{-1},
under industrially relevant conditions (pH 3 and 50 \%
CO\textsubscript{2}), indicated that the industrial SA-producing
strain\emph{S. cerevisiae} SUC632 was more robust than the congenic
laboratory strain CEN.PK113-7D, as the k\textsubscript{d} of the
industrial strain was about 3-fold lower than that of the laboratory
strain (Hakkaart et al., 2020). At near-zero growth rates in low-pH
retentostat cultures, k\textsubscript{d} values of the two strains were
not significantly different ( 0.0030 ± 0.0004 and 0.0030 ± 0.0006
h\textsuperscript{-1}, respectively) (Hakkaart et al., 2020).

In the ammonium-limited chemostat and retentostat cultures,
biomass-specific rates of SA production decreased asymptotically with
decreasing growth rate to an essentially constant value of 3 ± 0.1
mCmol/(g viable biomass)/h at specific growth rates below 0.02
h\textsuperscript{-1} (Figure 2B). This productivity was maintained
until, after 525 h (three weeks) the retentostat cultures were
terminated. These results demonstrated that cell retention in
ammonium-limited cultures can result in sustained, stable SA production
in the virtual absence of growth. The combined chemostat and retentostat
data showed that biomass-specific SA production rates increased linearly
with glucose consumption rate (Figure 7). This observation strongly
suggests that a coupling between the SA production rate and the flux
through primary metabolism, possibly as a consequence of altered
intracellular metabolite concentrations or altered expression of key
enzymes. Further research is required to elucidate the underlying
mechanisms and identify metabolic engineering targets for boosting SA
production rates in the absence of growth.

Use of membrane filters to achieve cell retention might be less feasible
on an industrial scale as blocking of effluent filters could prevent
long-time operation. Alternative means of cell retention could involve
use of fast-sedimenting yeast mutants (Oud et al., 2013) or
immobilization (Gulli et al., 2019; Nagarajan et al., 2014) might be
explored. Near zero growth conditions can also be achieved by
controlling the feed of a limiting nutrient in fed-batch processes. Due
to the absence of a liquid outflow, fed-batch cultivation results in
high final product titers, which are beneficial for the downstream
processing. Wahl (Wahl et al., 2017) studied growth of the same
SA-producing \emph{S. cerevisiae} strain in fed-batch cultures which,
during the final phase of cultivation, were starved for ammonium. In
contrast to our observation on nitrogen-limited retentostat cultures,
the specific SA production rate observed after ammonium depletion was
not stable but declined to a very low value within 90 h. As the authors
expressed specific SA productivity per total amount of biomass and did
not measure culture viability, this result may have reflected a loss of
culture viability cause by the low cultivation pH (Hakkaart et al.,
2020), complete nitrogen starvation and/or high SA concentrations. The
SA yield on glucose in the N-starved fed-batch cultures declined from
0.64 ± 0.01 to 0.53 ± 0.01 mol SA per mol glucose during the final phase
of fermentation (Wahl et al., 2017). In contrast, our ammonium-limited
retentostat cultures the SA yield on glucose stabilized to a value of
0.60 mol SA per mol glucose, which was maintained until the end of the
three-week retentostat runs (Figure 4). The lower SA yield in the
fed-batch cultures may be related to the much higher SA concentrations
(up to 0.6 mol/L, as compared to 63 ± 3 mmol/L in the retentostat
cultures). A high SA titer at a low cultivation pH will result in entry
of undissociated SA across the plasma membrane by passive diffusion,
followed by active export by the DCT02 transporter. Experiments
with\textsuperscript{13}C-SA demonstrated the relevance of such a futile
cycle, which necessitates an increased rate of glucose dissimilation to
CO\textsubscript{2} and water and, consequently, causes a lower SA yield
on glucose (Wahl et al., 2017).

Measurements of intracellular succinate and other TCA-cycle
intermediates in N-limited retentostat cultures showed levels consistent
with the genetic makeup of the industrial strain, whose metabolism has
been rewired towards reductive conversion of oxaloacetate to succinate
in the yeast cytosol by high-level expression of native and heterologous
enzymes (Jansen et al., 2013). Similar intracellular metabolite levels
were reported for ammonium-starved fed-batch cultures of the same strain
(Wahl et al., 2017). The very high mass action rations of the fumarase
reaction that were observed in both studies indicate a strong driving
force in the direction of fumarate synthesis and thus SA production.
However, \textsuperscript{13}C flux analysis in the fed-batch cultures
showed that the oxidative TCA cycle flux was significantly higher than
the reductive flux towards succinate. The net fumarase flux was
therefore in the direction of malate, which would require a \emph{in
vivo} mass action ratio below K\textsubscript{eq}. Wahl et al. explained
the high value of the mass action ratio from subcellular
compartmentation of malate (Wahl et al., 2017). The high malate to
fumarate ratio then suggests a possible limitation of cytosolic fumarase
for use of the reductive pathway towards SA, which could potentially be
relieved by modifying \emph{in vivo} kinetic and/or regulatory
characteristics of this key enzyme .

With this work we have shown that long-term extreme limitation by a
nutrient other than the carbon- and energy source enable stable
conversion of substrate into product in the virtual absence of growth.
Although we have used retentostat systems in this research, a similar
concept could be applied in repeated fed-batch cultivations, in which
the producing strain is used as a biocatalyst for prolonged periods of
time and conversion of substrate into biomass is minimized. Further
strain optimization should focus on improving glucose conversion rates
at near-zero growth rates and, in particular, on increasing cell
viability during prolonged cultivation at low pH.

\subsection*{Acknowledgements}

{\label{acknowledgements}}

This research was financed by the Netherlands BE-Basic research program.
We thank Dr. Alrik Los and Dr. Mickel Jansen at DSM for making the
industrial strain available for this study. We thank Cor Ras and
Patricia van Dam for analytical support.

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\subsection*{Figure legends}

{\label{figure-legends}}

\textbf{Figure 1} Culture viability (A,B), total and viable biomass
concentrations (C,D), total cell counts and average cell mass (E, F),
and specific growth and death rates (G,H) in aerobic, ammonium-limited
chemostat and retentostat cultures of \emph{S. cerevisiae} SUC632.
Cultures were grown on glucose, at pH 3 and were sparged with a 1:1
mixture of air and CO\textsubscript{2}. Left and right panels represent
data from steady-state chemostat cultures at different specific growth
rates and from retentostat cultures, respectively. In the retentostat
cultures, specific growth rates decreased over time (panel H). All data
represent averages ± standard errors of data from independent duplicate
cultures.

\textbf{Figure 2} Biomass specific consumption rates of glucose and
production rates of succinic acid and by-products as a function of the
specific growth rate (\selectlanguage{greek}µ\selectlanguage{english}) in aerobic, ammonium-limited chemostat and
retentostat cultures of \emph{S. cerevisiae} SUC632. Cultures were grown
on glucose, at pH 3 and were sparged with a 1:1 mixture of air and
CO\textsubscript{2}. q\textsubscript{byp} represents the sum of the
specific production rates of malate, ethanol, glycerol and acetate. All
data represent averages of results from duplicate chemostat and
retentostat cultures with their standard errors.

\textbf{Figure 3} Growth rate dependent division of consumed substrate
between growth, succinic acid and byproducts formation in aerobic,
ammonium-limited chemostat and retentostat cultures of \emph{S.
cerevisiae} SUC632, at different specific growth rates. Cultures were
grown on glucose, at pH 3 and were sparged with a 1:1 mixture of air and
CO\textsubscript{2}.

\textbf{Figure 4} Succinic acid yield on glucose during ammonium-limited
aerobic retentostat cultivation. Retentostat cultures were grown on
glucose, at pH 3 and were sparged with a 1:1 mixture of air and
CO\textsubscript{2}. Specific growth rates at the time points indicated
are shown in Figure 1H. All data represent averages with standard errors
obtained from duplicate retentostat cultures.

\textbf{Figure 5} Intracellular adenosine phosphate level, adenylate
energy charge and ATP/ADP ratio during aerobic, ammonium limited
retentostat cultivation of \emph{S. cerevisiae} SUC632. Retentostat
cultures were grown on glucose, at pH 3 and were sparged with a 1:1
mixture of air and CO\textsubscript{2}. Specific growth rates at the
time points indicated are shown in Figure 1H. Data represent the
averages and standard errors of measurements from duplicate cultures.

\textbf{Figure 6} Intracellular levels of the TCA cycle metabolites
(A-F) and mass action ratio's of fumarase and aconitase (G-H) during
aerobic, ammonium limited retentostat cultivation of SUC632. Retentostat
cultures were grown on glucose, at pH 3 and were sparged with a 1:1
mixture of air and CO\textsubscript{2}. Specific growth rates at the
time points indicated are shown in Figure 1H. Dashed blue lines in
Figures G and H represent the apparent \emph{in vivo} equilibrium
constants determined for\emph{S. cerevisiae} (Canelas et al., 2011).
Data represent the averages and standard errors of measurements from
duplicate cultures.

\textbf{Figure 7} Specific succinic acid production rate plotted as a
function of specific glucose consumption rate in aerobic,
ammonium-limited chemostat and retentostat cultures of \emph{S.
cerevisiae} SUC632. Cultures were grown on glucose, at pH 3 and were
sparged with a 1:1 mixture of air and CO\textsubscript{2}. All data
represent averages with standard errors obtained from duplicate
chemostat and retentostat cultivations.

\par\null

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