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\begin{document}

\title{A secretion-based dual fluorescence assay for high-throughput screening
of alcohol dehydrogenases}



\author[1]{Hongyuan Lu}%
\author[1]{Shiqin Yu}%
\author[1]{Fengyu Qin}%
\author[1]{Wenbo Ning}%
\author[2]{Xiaoqiang Ma}%
\author[1]{Kaiyuan Tian}%
\author[1]{Zhi Li}%
\author[1]{Kang Zhou}%
\affil[1]{National University of Singapore}%
\affil[2]{Singapore-MIT Alliance for Research and Technology Centre}%


\vspace{-1em}



  
  \date{June 10, 2024}


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\selectlanguage{english}
\begin{abstract}
Alcohol dehydrogenases (ADHs) play key roles in the production of
various chemical precursors that are essential in pharmaceutical and
fine chemical industries. To achieve practical application of ADHs in
industrial processes, tailoring enzyme properties through rational
design or directed evolution is often required. Here, we developed a
secretion-based dual fluorescence assay (SDFA) for high-throughput
screening of ADHs. In SDFA, an ADH of interest is fused to a mutated
superfolder green fluorescence protein (MsfGFP), which could result in
secretion of the fusion protein to culture broth. After a simple
centrifugation step to remove the cells, the supernatant can be directly
used to measure activity of the ADH based on a red fluorescence signal,
whose increase is coupled to formation of NADH (a redox co-factor of
ADHs) in the reaction. SDFA allows easy quantification of ADH
concentration based on the green fluorescence signal of MsfGFP. This
feature is useful in determining specific activity and may improve
screening accuracy. Out of five ADHs we have tested with SDFA, four ADHs
can be secreted and characterized. We successfully screened a
combinatorial library of an ADH from Pichia finlandica and identified a
variant with a 197-fold higher kcat/km value toward (S)-2-octanol
compared to its wild-type.%
\end{abstract}%



\sloppy


\section*{Keywords}

{\label{keywords}}

Protein secretion, Fluorescence assay, Alcohol dehydrogenase, Directed
protein evolution, Cascade reaction

\section*{1. Introduction}

{\label{introduction}}

Many drugs and value-added chemicals need to meet stringent
enantiopurity standards in pharmaceutical and fine chemical industries.
During the last decade, there has been a growing interest in using
enzymes for producing chiral chemicals. This is mainly driven by the
inherent advantages of biocatalysis over chemical synthesis, such as
high chemo-, regio- and enantio-selectivities, mild reaction conditions
and using less toxic reagents or solvents (Nealon, Musa, Patel, \&
Phillips, 2015; Zheng et al., 2017).

In particular, alcohol dehydrogenases (E.C. 1.1.1.x; x=1 or 2, ADHs) are
among the most investigated redox enzymes. ADHs are a group of
NAD(P)(H)-dependent enzymes that catalyze the interconversion between
alcohols and ketones or~aldehydes. Because of their strict
stereoselectivity, ADHs have been employed in the production of various
chiral products with high optical purity (Chen, Liu, Lin, \& Zheng,
2016; Kara et al., 2014; Savile, Gruber, Mundorff, Huisman, \& Collier,
2014). However, the performance of wild-type ADHs is rarely adequate for
industrial processes. In general, tailoring enzyme properties, such as
catalytic activity, stereoselectivity, substrate specificity,
thermostability and tolerance of organic solvent is often required to
realize the practical application of ADHs (R. Zhang, Xu, \& Xiao, 2015).

Directed evolution is one of the most effective methods to engineer
ADHs. An ADH may be randomly mutated by using error-prone PCR to
identify beneficial mutations. Putative critical residues of an ADH may
also be identified based on docking the substrate/product into
experimental-determined or predicted tertiary structure of the ADH.
Beneficial mutations at these putative residues would be characterized
through mutagenesis. Often, multiple beneficial mutations need to be
combined and the aforementioned process may be iterated.

In directed evolution, screening the mutants is usually the
rate-limiting unit operation, because the rate of hitting a beneficial
mutation is low. The assay commonly used to quantify activity of ADH is
to monitor the production or depletion of NAD(P)H based on absorbance at
340 nm as these enzymes couple their enzymatic reactions with the
nicotinamide cofactors (Forrest \& Gonzalez, 2000). Usually cells
expressing an ADH was lysed prior to the assay. Unfortunately, this
approach is not suitable for high-throughput screening, because of the
strong absorption of UV light by cell lysates (Mayer \& Arnold, 2002).

Alternatively, colorimetric assays that combine the redox reaction with
a dye-forming reaction have been developed to reduce the noise, because
the absorption of visible light by cell lysates was weaker than that of
UV light. One of the most widely adopted colorimetric approaches is the
nitroblue tetrazolium/phenazine methosulfate (NBT/PMS) assay (Fibla \&
Gonz\selectlanguage{ngerman}àlez-Duarte, 1993). In the presence of the transferring agent PMS,
NBT can be reduced by the NAD(P)H regenerated by an ADH-catalyzed
reaction to produce an insoluble blue-purple formazan dye, whose
absorbance can be measured at 580 nm. This colorimetric assay has been
used in high-throughput screening to identify novel or improved ADHs,
such as 6-phosphogluconate dehydrogenase (Huang, Chen, Zhong, Kim, \&
Zhang, 2016), glycerol dehydrogenase (H. Zhang, Lountos, Ching, \&
Jiang, 2010), halohydrin dehydrogenase (Fox et al., 2007), and
phenylacetaldehyde reductase (Makino, Dairi, \& Itoh, 2007). However,
this colorimetric assay still needs improvement due to its background
noise and limited detection range. Moreover, this assay also fails to
enable a continuous measurement of ADHs activity as the final colour
product is insoluble, resulting in precipitation that interferes with
the absorbance quantification.

In addition, the aforementioned assays generally suffer from two common
issues when they are used to screen cells that express enzyme. First, in
order to release target enzyme from host cells for determining catalytic
activity, cell lysis is often required by these methods. However, once
the cells are lysed, together with the enzyme of interest, all cytosolic
components are simultaneously released, resulting in a complex matrix
that could reduce the sensitivity and accuracy of the assay implemented
(Contreras-Llano \& Tan, 2018; Mayer \& Arnold, 2002). For example,
cytosolic components, such as nicotinamide cofactor, autofluorescent or
chromogenic metabolites may miscount for the catalytic activity of the
target enzyme. Non-specific interactions with endogenous enzymes (e.g.
endogenous ADHs) could also interfere with the target enzyme reaction if
they share similar substrate specificity.

Furthermore, when these assays were applied in high-throughput format,
such as microtiter plate screening, the catalytic activity was usually
evaluated without normalizing by the amount of enzyme interrogated.
Because, it is time-consuming and tedious to quantify the enzyme amount
for each variant in a large-scale screening effort. Consequently, this
inevitably leads to an unfair comparison among the mutants. The quantity
of the target enzyme in each mutant sample could vary due to fluctuating
protein expression level and nonhomogeneous cell growth rate. Overall,
this may not only produce a considerable number of false positives, but
also miss out some of the true positives. It will be desirable to have a
simple and easy way to quantify the enzyme amount for each mutant so
that their catalytic activity can be normalized for a fairer screening.

To tackle these limitations in this study, we developed a
secretion-based dual fluorescence assay (SDFA) for high-throughput
screening of mutated ADHs. This assay is based on a cascade reaction
leading to the formation of a red fluorescent molecule. It also uses a
protein secretion system translocating the target enzyme to a simpler
reaction matrix than cell lysate.

As shown in Figure 1, in the presence of an enzyme (diaphorase),
resazurin reacts with the NAD(P)H generated from the ADHs-catalysed
reaction, leading to the generation of the highly fluorescent
molecule---resorufin---whose emission signal can be easily detected at
588 nm when excited at 535 nm. Since the emission falls within the range
of red light, in which fewer \emph{Escherichiacoli} endogenous molecules
emit fluorescence, there is less interference from the reaction mixture
and spent culture media.

Meanwhile, we combined the cascade reaction with a protein secretion
system which was enabled by a mutated superfolder green fluorescent
protein (MsfGFP) (Z. Zhang et al., 2017). By fusing ADHs to MsfGFP, the
fusion proteins can be efficiently secreted out of the host cells for
enzymatic activity quantification. Therefore, cell lysis was not
required by this assay. This led to a much simpler reaction matrix that
was free of complex cytosolic components. As a result, the interference
caused by intracellular metabolites and endogenous enzymes was
eliminated. Also, this secretion system simplified the screening
procedure and avoided the potential inconsistency of lysis efficiency.

Additionally, another merit of this assay is that the green fluorescence
of the MsfGFP fusions was linearly proportional to its protein amount.
This provides a simple and efficient way to quantitate the amount of
target enzyme in the reaction matrix. Therefore, normalization of the
catalytic activity of different mutants can be easily achieved, further
improving the accuracy of the assay.

In this study, we have collected experimental data to support the
claimed advantages of SDFA, and successfully demonstrated the usefulness
of SDFA through engineering of an ADH, which led to substantial
improvement of its catalytic efficiency.

\section*{2. Materials and Methods}

{\label{materials-and-methods}}

\subsection*{Materials}

{\label{materials}}

All the chemicals and diaphorase used in this study were purchased from
Sigma-Aldrich Pte. Ltd., Singapore, unless otherwise stated. The QIAprep
Spin Miniprep kit for plasmid extraction was purchased from Qiagen Pte.
Ltd., Singapore. The GeneJET gel extraction kit for gel extraction was
purchased from Thermo Fisher Scientific, Singapore.

\subsection*{Plasmid and strain
construction}

{\label{plasmid-and-strain-construction}}

The MsfGFP gene sequence was kindly provided by Dr. Ma Lixin from Hubei
University, Wuhan, People's Republic of China. Synthetic ADHs genes and
all the used oligonucleotides were ordered from Integrated DNA
Technologies. Plasmid construction was carried out based on the
Guanine--Thymine standard (Ma et al., 2019). The fusion proteins consist
of an N-terminal MsfGFP or eGFP which is separated by a linker with an
TEV Protease recognition site sequence (ENLYFQG) from an ADH. The DNA
sequence of all the constructed plasmids was verified by Sanger
sequencing. A full list of plasmids constructed in this study is
provided in Table S1. \emph{E coli} DH5\selectlanguage{greek}α \selectlanguage{english}(C2987H, New England Biolabs)
was used as cloning host and \emph{E. coli} BL21 (C2527H, New England
Biolabs) was used for protein expression. Plasmid transformation was
carried out by using the heat-shock method provided in the
manufacturer's instruction.

\subsection*{Protein expression and
purification}

{\label{protein-expression-and-purification}}

A single colony of an \emph{E. coli} BL21 strain harboring one of the
expression plasmids in Table S1 was inoculated into 3 ml of Luria
Bertani broth (LB) with 50 \selectlanguage{greek}µ\selectlanguage{english}g/ml spectinomycin and grown overnight at 37
\selectlanguage{ngerman}°C/250 rpm. The overnight-grown culture (using 500 \selectlanguage{greek}µ\selectlanguage{english}l) was scaled up
with 300-fold dilution in a 500-ml shake flask and grown to a cell
density of 0.6 (optical density {[}OD600{]}) at 37 \selectlanguage{ngerman}°C/250 rpm. Protein
expression was induced by adding 0.1 mM of isopropyl
\selectlanguage{greek}β-\selectlanguage{english}D-1-thiogalactopyranoside (IPTG) and cells were cultured for 22 hours
at 30 \selectlanguage{ngerman}°C/250 rpm.

For isolation of the intracellularly expressed (\emph{R} )-2-octanol
dehydrogenase (PfODH), cells were harvested by centrifugation at 4,000 g
for 10 mins. Cell pellets were then resuspended in equilibrium buffer
(20 mM sodium phosphate, 300 mM sodium chloride, and 10 mM imidazole; pH
7.4), and mechanically lysed by a cell disrupter (OS,
Constant~Systems~Ltd.). The resulting cell lysate was centrifuged at
10,000 g for 20 mins to obtain the supernatant that contains soluble
proteins. For isolation of the extracellularly expressed MsfGFP fusion
proteins, the induced cell culture was centrifuged at 4,000 g for 20
mins to obtain the supernatant that accommodates secretory proteins.
Target proteins from the above two types of supernatant were both
purified by HisPur\selectlanguage{english} Ni-NTA Resin (Thermo Fisher Scientific) according to
the manufacturer's instruction. Desalting of the protein elution was
carried out by using Sephadex G-25 PD-10 columns (GE Healthcare)
according to the manufacturer's instruction. The concentration of
purified proteins was determined by the Micro BCA\selectlanguage{english} Protein Assay kit
(Thermo Fisher Scientific). The absorbance of assay mixtures was
measured by the Infinite 200\textsuperscript{(r)} PRO microplate plate
reader~(Tecan Group AG). The presence of the purified target proteins
was verified by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) using
Mini-PROTEAN TGX Precast Protein Gels (4561083, Bio-Rad). Gel image was
taken by the Gel Doc EZ Gel Documentation System (Bio-Rad).

\subsection*{Fluorescence observation and SDS-PAGE analysis of
supernatant
samples}

{\label{fluorescence-observation-and-sds-page-analysis-of-supernatant-samples}}

The protein expression of the MsfGFP/eGFP-ADH fusion proteins was
induced as described above. The grown cell cultures were then
centrifuged at 4,000 g for 20 mins to obtain the supernatants. To
observe the fluorescence of the fusion proteins, the supernatant samples
were excited by blue light using a Safe Imager\selectlanguage{english} 2.0 Blue Light
Transilluminator (Thermo) and the image was taken through an amber
filter unit. Afterwards, the supernatant samples were concentrated
5-fold by Microcon(r) centrifugal filter devices (Merck Millipore) for
SDS-PAGE analysis.

\subsection*{The coupled enzyme assay with purified ADH/secreted fusion
proteins}

{\label{the-coupled-enzyme-assay-with-purified-adhsecreted-fusion-proteins}}

The coupled enzyme assay reagent was prepared as a mixture consisting of
4 U/ml diaphorase, 1 mM resazurin, 1 mM substrate (for ADH), 1 mM
NAD\textsuperscript{+} and 0.1 M Tris buffer (pH 8.0). To examine the
catalytic activity of PfODH toward various substrates, the purified
PfODH was first diluted with 0.1 M Tris buffer to yield a final
concentration of 20 \selectlanguage{greek}μ\selectlanguage{english}g/ml. Subsequently, 50 \selectlanguage{greek}µ\selectlanguage{english}l of each diluted PfODH
sample was then pipetted into the wells of a 96-well back polystyrene
microtiter plate (Costar 3603, Corning). Assay reactions were initiated
by adding 50 \selectlanguage{greek}µ\selectlanguage{english}l of the assay mixture to each sample well, immediately
followed by a continuous measurement of the fluorescence of resorufin
(Ex/Em=535/588 nm) with the ``Kinetic Measurement'' mode in i-control
software for 10 mins by the Infinite 200\selectlanguage{ngerman}\textsuperscript{®} PRO
microplate plate reader~(Tecan Group AG).

The same procedure was used for determining the catalytic activity of
secreted fusion proteins. Cultures of the MsfGFP-fused ADHs were
centrifuged at 4,000 g for 20 mins to obtain the supernatant that
accommodates secretory proteins. 50 \selectlanguage{greek}µ\selectlanguage{english}l of the supernatant sample was
used for the assay. Similarly, for high throughput screening experiment,
induced cell cultures in 96 deep well plates (Greiner Bio-One GmbH) were
centrifuged at 4,000 g for 20 mins. 50 \selectlanguage{greek}µ\selectlanguage{english}l of the supernatant sample from
each mutant was transferred to a well in a 96-well microtiter plate, and
the fluorescence of MsfGFP-fused mutants (Ex/Em=488/525 nm) was
determined first by plate reader. Subsequently, 50 \selectlanguage{greek}µ\selectlanguage{english}l of the assay
mixture was added to the well, immediately followed by a continuous
measurement of the fluorescence of resorufin (Ex/Em=535/588 nm) by plate
reader.

\subsection*{Kinetic analysis}

{\label{kinetic-analysis}}

Kinetic analysis of the wild-type PfODH and the selected PfODH variant
was performed by determining the production of NADH (\selectlanguage{greek}λ \selectlanguage{english}= 340 nm) through
absorbance measurement. Assays were carried out in 200 \selectlanguage{greek}μ\selectlanguage{english}l of PBS buffer
(100 mM, pH 7.5) containing 6 mM NAD\textsuperscript{+}, at room
temperature, using 13.3 \selectlanguage{greek}μ\selectlanguage{english}M of the wild-type PfODH or 0.055 \selectlanguage{greek}μ\selectlanguage{english}M of the
selected PfODH variant at substrate concentrations ranging from 0.125 mM
to 5.0 mM. The extinction coefficient used for NADH under the assay
conditions was 6220 M\textsuperscript{-1}cm\textsuperscript{-1}. All
assays were performed in triplicate. The enzyme kinetic parameters
k\textsubscript{m}, and k\textsubscript{cat} were calculated from the
enzyme progress curves via the Lineweaver-Burk plot.

\section*{3. Results and discussion}

{\label{results-and-discussion}}

\subsection*{A cascade reaction enables fluorogenic determination of ADH
activity}

{\label{a-cascade-reaction-enables-fluorogenic-determination-of-adh-activity}}

As shown in Figure 1, the catalytic activity of ADHs in an oxidation
reaction can be expressed as the rate of forming NAD(P)H. Although it is
possible to quantitate NAD(P)H by directly measuring its light
absorbance (UV 340 nm), this method suffers from low sensitivity and
high background noise (R. Zhang et al., 2015). Alternatively, measuring
the fluorescence of NAD(P)H Ex/Em=340/445 nm) can be used to quantify
NAD(P)H and has shown a greater sensitivity compared to the
absorbance-based approach (Held, 2007). However, our experiments showed
that although the linear correlation between NADH concentration and its
fluorescence can be obtained when NADH was serially diluted with
deionized water, buffer or fresh media (Figure S1 A-D), such a
correlation cannot be established when spent media was used as the
diluent (Figure S1 E-F). These results suggest that metabolites released
by growing cells can substantially interfere with the fluorescence
detection of NADH, and therefore such fluorescence assay is not suitable
for determining the NADH level in crude samples such as cell lysate or
culture supernatant.

To tackle this issue, we employed an indicator enzymatic reaction in
which NADH was used by diaphorase to reduce resazurin into resorufin,
which is highly fluorescent (Figure 1). The main advantage of this
system is that the long excitation/emission wavelengths of resorufin
(Ex/Em=535/588 nm) can reduce the interferences from cellular
metabolites, because most fluorescent molecules in cell lysates are
excited in the UV range (Simeonov \& Davis, 2004). As expected, our
results showed that fluorescence of the formed resorufin responded
linearly to NADH concentration across different medium backgrounds,
including spent media (Figure S1 G-L). Meanwhile, this reaction system
offers a much more extended linear detection range (0.1-700 \selectlanguage{greek}μ\selectlanguage{english}M) compared
to NADH fluorescence-based approach (0.1-200 \selectlanguage{greek}μ\selectlanguage{english}M when the medium
background is deionized water). Additionally, through transforming the
unstable NADH into the more stable resorufin, this reaction system
provides a more durable and accurate fluorescent signal, allowing a
reliable and continuous fluorogenic determination of NADH concentration.

Given these advantages, we may couple an ADH-catalysed reaction with
this indicator system to form a coupled enzyme assay for the ADH (Figure
1). To examine whether the ADH activity can be determined by this
cascade reaction, we employed an enantioselective ADH, (\emph{R}
)-2-octanol dehydrogenase (PfODH) as the model enzyme. In the cascade
reaction mixture, the purified enzyme PfODH was supplied with cofactor
NAD\textsuperscript{+} and substrate to initiate target reactions.
Concurrently, excessive amount of resazurin and diaphorase were provided
in the mixture to stoichiometrically convert NADH into fluorescent
resorufin. The enzymatic activity of ADH was thus proportional to the
rate at the red fluorescence increased. As shown in Figure 2, the
catalytic activity of PfODH toward various substrates, including its
native substrate ((\emph{R} )-2-octanol) and other non-native substrates
(4-Fluoro-\selectlanguage{greek}α-\selectlanguage{english}methylbenzyl alcohol, 1-(3-Methylphenyl)ethanol and
1-Phenylethanol), were determined by following the respective increases
in fluorescence of resorufin over time using a plate reader. In
contrast, the negative controls without adding PfODH all exhibited no
fluorescence increase over time. These results showcased that the
coupled enzyme assay can enable a sensitive and continuous,
time-dependent monitoring of the ADH-catalysed reaction.

\subsection*{Incorporating the coupled enzyme assay with a protein
secretion
system}

{\label{incorporating-the-coupled-enzyme-assay-with-a-protein-secretion-system}}

High-throughput screening used in enzyme evolution has relied heavily on
the use of cell-based approach (Zeymer \& Hilvert, 2018). However, there
are two major defects with this method: (i) It requires cell lysis that
could introduce interference to the screening assay; (ii) It relies on
the false assumption that the amount of enzyme being assayed is uniform
across all mutants, therefore resulting in an unfair comparison that
consequently increases false positive or negative rates. To further
develop our coupled enzyme assay into a better screening method, we set
out to address these two limitations.

In particular, we proposed to combine the cascade reaction with a
protein secretion system which is based on MsfGFP. The addition of such
a secretion system can potentially offer two unique features to our
assay. First, it has been reported that when recombinant proteins were
fused with MsfGFP, it can lead to the auto-secretion of the fusion
protein without disturbing their conformation and function (Z. Zhang et
al., 2017). Such a non-specific auto-secretion system is highly
desirable because cell lysis can therefore be exempted, and the secreted
target enzyme can be assayed in the culture broth, which is a much
simpler matrix than cell lysate. Second, MsfGFP can serve as a protein
expression reporter for its passenger protein. It has been demonstrated
that the fluorescence of sfGFP (non-mutated superfolder GFP) fusion
protein is linearly proportional to the protein quantity (P\selectlanguage{ngerman}édelacq,
Cabantous, Tran, Terwilliger, \& Waldo, 2006). This feature can
potentially be used to normalize the enzyme activity of each mutant
during high throughput screening, since the fluorescence of MsfGFP
fusion protein can be easily and efficiently measured using fluorescence
plate reader.

To evaluate whether the MsfGFP-guided secretion system is generally
applicable to ADHs, we fused the MsfGFP or eGFP (enhanced GFP, a control
which would not lead to protein secretion) to N-terminus of five ADHs,
including PfODH from \emph{Pichia finlandica,} (\emph{S}
)-1,3-butanediol dehydrogenase (CpSADH) from \emph{Candida
parapsilosis,} glycerol dehydrogenase (GlyDH) from \emph{E.} \emph{coli}
, Meso-2,3-butanediol dehydrogenase (BDH\_KP) from \emph{Klebsiella
pneumoniae} and 2,3-butanediol dehydrogenase (BDH\_BS) from
\emph{Bacillus subtilis} . The protein expression of the MsfGFP/eGFP-ADH
fusion proteins was induced by IPTG. The grown cell cultures were then
centrifuged to obtain the supernatants for examining whether the fusion
proteins were secreted out of the cells. As shown in Figure 3, all the
supernatants of the cells expressing the MsfGFP fusion proteins
exhibited fluorescence to various degrees, whereas all the controls did
not show any fluorescence. Meanwhile, the presence of secreted MsfGFP
fusion proteins and the absence of eGFP fusion proteins in supernatant
were further confirmed by SDS-PAGE analysis (Figure 3). Consistent with
the fluorescence observation, the respective protein bands of MsfGFP
fusion proteins showed varying degrees of staining, further indicating
different levels of secretion for the MsfGFP fusion proteins. These
results suggest that secretion of five ADHs can be realized by using
this MsfGFP-guided auto-secretion system, although with various degree
of secretion efficiency.

After we confirmed MsfGFP can enable the secretion of ADHs, we moved
forward to examine whether the secreted ADHs' activity can be determined
by the coupled enzymatic assay. Assay reaction components were directly
added to the culture supernatants of each fusion protein. As shown in
Figure 4, the supernatant samples of most MsfGFP-guided ADHs (GlyDH,
CpSADH, PfODH, and BDH\_KP) generated various fluorescence increases
over time through the assay reaction, whereas there was no fluorescence
change for all the supernatant sample of eGFP fusion proteins. However,
unlike other ADHs, the rate of red fluorescence increase was low for
MsfGFP-BDH\_BS, which may be mainly due to its low secretion level.
Nonetheless, these results demonstrated that the MsfGFP-guided secretion
system can be combined with the coupled enzyme assay to establish a
secretion-based assay for determining activity of some ADHs.

\subsection*{Normalizing ADH activity by the fluorescence of
MsfGFP}

{\label{normalizing-adh-activity-by-the-fluorescence-of-msfgfp}}

We next assessed the feasibility of using the fluorescence of MsfGFP to
normalize the catalytic activity of ADHs among different mutants in
high-throughput screening. To mimic the actual screening condition of
the secreted ADHs, we purified the fusion protein MsfGFP-PfODH and
serially diluted it into different concentrations with cell-free spent
medium. The green fluorescence (Ex/Em=488/525 nm) of the diluted enzyme
samples were first determined by fluorescence plate reader. The green
fluorescence intensity exhibits a strong linear correlation to the
concentration of MsfGFP-PfODH (Figure S2A). Subsequently, the catalytic
activity of the MsfGFP-PfODH samples with various concentrations were
determined by measuring their respective red fluorescence (Ex/Em=488/525
nm) after adding coupled enzyme assay reagents. When (\emph{R}
)-2-octanol was used as the substrate, the samples with higher
concentrations of MsfGFP-PfODH exhibited higher red fluorescence
intensities (Figure 5). Apparently, without considering the different
amounts of enzyme in the assay, the specific activity of these enzyme
samples, which was supposed to be the same, can be misrepresented based
on the red fluorescence measurement. To address this limitation, we used
the green fluorescence intensity, which reflected the total amount of
the ADH in the assay, to normalize the red fluorescence intensity of
each sample. The normalized signal of each sample was very close to each
other (Figure 5), indicating similar specific activity. Moreover, Figure
S3 showed that the same normalizing effect of MsfGFP's fluorescence can
also be applied when a different substrate ((\emph{R} )-2-butanol) was
used in the assay reaction. In addition, our further experiments also
proved that this normalizing feature of MsfGFP also worked for another
ADH (Figure S4). These results demonstrated that the apparent activity
of ADHs can be normalized by the fluorescence of MsfGFP fusion proteins
to enable a fairer comparison of enzyme mutants, thus reducing the false
positive and false negative rates during screening.

\subsection*{Using the secretion-based dual fluorescent assay in protein
directed
evolution}

{\label{using-the-secretion-based-dual-fluorescent-assay-in-protein-directed-evolution}}

Considering the aforementioned advantages, we combined the
resorufin-based cascade reaction system and the MsfGFP-guided secretion
system to establish the secretion-based dual fluorescent assay (SDFA)
for directed evolution of ADHs. On the one hand, the cascade reaction
system offers a sensitive and continuous determination of ADHs activity
through a red fluorescence measurement. On the other hand, MsfGFP-guided
secretion system provides an easy and efficient way to translocate the
target ADHs to a simpler reaction matrix and to normalize the enzyme's
activity through a green fluorescence measurement.

In the workflow of SDFA (Figure 6), the mutated genes of an ADH were
first fused to the C-terminus of \emph{msfgfp} to create a plasmid
library. The host cells (\emph{E. coli} ) were then transformed with the
constructed plasmids and were spread on selection agar plates. Fully
grown mutant colonies were inoculated in liquid culture medium and
induced for enzyme expression. Subsequently, the cell cultures were
centrifuged down to obtain the supernatants that contained the secreted
enzymes for further enzyme screening. The green fluorescence of the
supernatants from the enzyme mutants was first determined by a plate
reader. Afterwards, assay reagents were added to the supernatant samples
to monitor the red fluorescence. Finally, the catalytic activity of each
mutant was normalized (red/green fluorescence) to identify the true
positive mutant hits.

As a case study for demonstrating the applicability of SDFA in
high-throughput screening, we employed PfODH as the model enzyme and
sought to engineer its stereoselectivity. In oil industry, alcohol
oxidation is one of the most extensively studied reactions as it plays a
key role in converting petroleum-based feedstock into essential building
blocks for a wide variety of chemicals (Parmeggiani, Matassini, \&
Cardona, 2017). PfODH belongs to the short chain dehydrogenase/reductase
(SDR) superfamily. It has a very broad substrate specificity and can
catalyse the oxidation of alcohols to the corresponding aldehydes or
ketones, which are the ubiquitous chemical precursors in pharmaceutical
and fine chemical industries (H. Yamamoto \& Kudoh, 2013). However,
PfODH has a substrate preference toward\emph{R} -enantiomer of alcohols,
thus resulting in incomplete conversion of racemic alcohol, i.e.
\emph{S} -enantiomer of alcohols cannot be fully utilized (H. Yamamoto
\& Kudoh, 2013). From an~atom economy point of view, enzymes that can
oxidize both enantiomers of alcohol are more desirable for this
application.

Therefore, we set out to engineer the stereoselectivity of PfODH through
screening enzyme variants with enhanced activity toward\emph{S}
-enantiomer of alcohol. To achieve this goal, we constructed a
combinatorial library, PfODH-G99VWA-H150WTA-Y193RBT (containing 60
genotypes), on critical sites G99, H150, and Y193 (Figure 7), which are
involved in the formation of the active cavity and extend their side
chains into the cavity of PfODH. These corresponding sites were
suggested as the switch to tune the stereoselectivity of the enzymes in
the superfamily of SDRs (Qin et al., 2018). Variation of steric
hindrance and polarity on sites of G99, H150, and Y193 might influence
the stereo-preference of PfODH (Qin et al., 2018). Therefore, the
mentioned library was constructed as the demonstration to test the
applicability of SDFA as a high throughput screening method in directed
evolution of ADHs.

To ensure full coverage of the design space for our constructed library,
307 clones were picked for enzyme expression. These 307 mutants were
assayed by SDFA in 96 well plate format (Figure S5). Through screening,
top 10 variants that exhibited the highest catalytic activity against
(\emph{S} )-2-octanol were selected for sequencing to identify their
genome type. Sequence analysis revealed that 8 out of the 10 selected
mutants were identified as PfODH\textsuperscript{G99L/A155L/Y193A},
whereas the remaining two mutants are
PfODH\textsuperscript{G99Q/A155L/Y193A} and
PfODH\textsuperscript{G99V/A155L/Y193A}. Given the fact that
PfODH\textsuperscript{G99L/A155L/Y193A} was dominant in the top 10
positive hits and also showed higher catalytic activity than the other
two variants, PfODH\textsuperscript{G99V/A155L/Y193A} was identified as
the best mutant from this library screening. To cross-validate the
screening result of SDFA, PfODH\textsuperscript{G99L/A155L/Y193A} was
expressed in shaking flask cultures and purified for further determining
its kinetic parameter. Our results showed that
PfODH\textsuperscript{G99L/A155L/Y193A} had a 197-fold higher
k\textsubscript{cat}/k\textsubscript{m} value toward (\emph{S}
)-2-octanol in comparison with wild-type PfODH. These results
demonstrated the capacity of SDFA to identify improved enzyme variants
from a library of mutants, illustrating the usefulness of this assay in
high-throughput screening of ADH mutants.

\section*{4. Conclusion}

{\label{conclusion}}

In summary, we report here the development of a sensitive, simple, and
efficient fluorogenic assay that is suitable for high-throughput
screening of ADHs. We demonstrated the potential of SDFA by performing
directed evolution on PfODH through screening a combinatorial library.
The selected enzyme variant exhibited a substantially higher
k\textsubscript{cat}/k\textsubscript{m} value toward (\emph{S}
)-2-octanol compared to its wild-type. In addition, the secretion-based
nature of SDFA exempts the need for cell lysis, thus bypassing its
disadvantages including the release of interfering cytosolic components,
the additional cost of lysis reagent, the potential inconsistency of
lysis efficiency, the extra effort of preserving intact culture for
genome recovery. From this perspective, SDFA not only provides a
sensitive and reliable way for determining ADHs activity, but also
simplifies the screening procedure.

To the best of our knowledge, SDFA is the first secretion-based assay
that can easily enable normalizing the catalytic activity of ADHs for
high throughput screening. SDFA is in theory also applicable to other
NAD(P)(H)-dependent enzymes besides ADHs and is suitable for
high-throughput screening methods such as 96/384 well plate format.
Although the method we have developed so far only works with the
oxidative direction of the ADH-catalyzed reaction, SDFA should still be
attractive to many protein engineers, because there are a large number
of useful oxidative reactions catalyzed ADHs.

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\section*{Figure legend:}

{\label{figure-legend}}

\section*{}

{\label{section}}

\textbf{Figure 1} Working principle of SDFA. Target ADH is translocated
to the extracellular environment by a MsfGFP-guided secretion system.
Subsequently, the enzyme activity of ADH is determined by a cascade
reaction in which NADH produced by the ADH-catalyzed reaction was used
by diaphorase to reduce resazurin into resorufin, which is highly
fluorescent. When excessive amount of resazurin and diaphorase were
provided, the rate of NADH formation was proportional to the rate at
which the red fluorescence increased.

\textbf{Figure 2.} Catalytic activity of PfODH toward various substrates
determined by the coupled enzyme assay. The increase of resorufin
fluorescence (Ex/Em=535/588 nm) was monitored for the PfODH-catalyzed
reactions with \textbf{(A)} (\emph{R} )-2-octanol,
\textbf{(B)}1-(3-Methylphenyl)ethanol, \textbf{(C)}
4-Fluoro-\selectlanguage{greek}α-\selectlanguage{english}methylbenzyl alcohol, and \textbf{(D)} 1-Phenylethanol
respectively. All assays were performed in a final reaction volume of
100 \selectlanguage{greek}μ\selectlanguage{english}l with the following conditions: 10 \selectlanguage{greek}µ\selectlanguage{english}g/ml purified PfODH, 2 U/ml
diaphorase, 0.5 mM resazurin, 0.5 mM alcoholic substrate, 0.5 mM
NAD\textsuperscript{+}, 0.1 M Tris solution (pH 8.0). Assay reactions
without adding purified PfODH served as the controls for each substrate
reaction. RFU: relative fluorescence units.

\textbf{Figure 3} . Fluorescence image of the supernatants of the cells
expressing the MsfGFP/eGFP fusion proteins or eGFP (upper part of the
figure). The supernatant samples were excited by blue light (Safe
Imager 2.0 Blue Light Transilluminator, Thermo) and the image was taken
through an amber filter unit. SDS-PAGE analysis of the concentrated
supernatant samples of the cells expressing the MsfGFP/eGFP fusion
proteins or eGFP (lower part of the figure).

\textbf{Figure 4.} Catalytic activity of the secreted MsfGFP fusion
proteins determined by the coupled enzyme assay. The increase of
resorufin fluorescence (Ex/Em=535/588 nm) was monitored for the
reactions catalyzed by the secreted MsfGFP fusion proteins.
\textbf{(A)}MsfGFP-GlyDH and eGFP-GlyDH were characterized by using
1,3-butanediol as the substrate. \textbf{(B)} MsfGFP-CpSADH and
eGFP-CpSADH were characterized by using (\emph{S} )-2-octanol as the
substrate.\textbf{(C)} MsfGFP-PfODH and eGFP-PfODH were characterized by
using (\emph{R} )-2-octanol as the substrate. \textbf{(D)}
MsfGFP-BDH\_KP and eGFP-BDH\_KP were characterized by using
Meso-2,3-butanediol as the substrate. The assay was carried out by
mixing the supernatant sample and the assay reagent mixture at
volumetric ratio of 1:1. All assays were performed in a final reaction
volume of 100 \selectlanguage{greek}μ\selectlanguage{english}l with the following conditions: 50 \selectlanguage{greek}μ\selectlanguage{english}l supernatant
sample, 2 U/ml diaphorase, 0.5 mM resazurin, 0.5 mM alcoholic substrate,
0.5 mM NAD\textsuperscript{+}, 0.05 M Tris solution (pH 8.0). RFU:
relative fluorescence units.

\textbf{Figure 5.} Catalytic activity of ADHs normalized by the
fluorescence of MsfGFP. Reactions catalyzed by various concentrations of
MsfGFP-PfODH with substrate (\emph{R} )-2-octanol. Red fluorescence
represents the fluorescence intensity of resorufin (Ex/Em=535/588 nm)
recorded at a time point within the linear part of the progress curve of
MsfGFP-PfODH catalyzed reactions. The same time point measurement was
taken for all the reactions with various enzyme concentrations. Green
fluorescence represents the fluorescence intensity of MsfGFP-PfODH. The
normalized signal is calculated by dividing the red fluorescence signal
by the green fluorescence signal. The assay was carried out by mixing
the MsfGFP-PfODH sample and the assay reagent mixture at volumetric
ratio of 1:1. All assays were performed in a final reaction volume of
100 \selectlanguage{greek}μ\selectlanguage{english}l with the following conditions: 50 \selectlanguage{greek}μ\selectlanguage{english}l enzyme sample, 2 U/ml
diaphorase, 0.5 mM resazurin, 0.5 mM alcoholic substrate, 0.5 mM
NAD\textsuperscript{+}, 0.05 M Tris solution (pH 8.0). Each bar
indicates the mean value of three biological replicates and the
corresponding error bar indicates the standard deviation. RFU: relative
fluorescence units.

\textbf{Figure 6.} Schematic overview of the SDFA workflow for the
directed evolution of ADHs. 1-2: Mutated ADHs' genes were fused to the
C-terminus of MsfGFP to create a plasmid library. 3-4: The host cells
were transformed with the constructed plasmids and were spread on
selection agar plates. 5: Grown mutant colonies were inoculated in
liquid medium and induced for enzyme expression. 6: The cell cultures
were centrifuged down to obtain the supernatants that contained the
secreted enzymes. The green fluorescence of the supernatants from the
enzyme mutants was determined. 7: Assay reagents were added to the
supernatant samples to monitor the red fluorescence. 8: The catalytic
activity of each mutant was normalized to identify positive hits.

\textbf{Figure 7.} The predicted conformations of (R)-2-octanol (cyan
ball-stick) and NAD\textsuperscript{+} (yellow stick) in the active site
of PfODH. The catalytic residues (S148 and Y161) are shown in blue
sticks. The critical residues (G99, H150, and Y193), which affect the
substrate specificity and enantioselectivity, are shown in green sticks.
The substrate, (\emph{R} )-2-octanol, was built by the builder tool of
PyMol (DeLano Scientific) in productive conformation. Model structure of
PfODH--NAD\textsuperscript{+} complex was constructed based on homology
modeling method using Swiss-Model server
(\url{https://swissmodel.expasy.org}). The structure of glucose
dehydrogenase from \emph{Bacillus megaterium} IWG3 (K. Yamamoto et al.,
2001), BmGDH (PDB: 1GCO), was selected as the template. They share 40\%
sequence identity.

\par\null\par\null

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