A Cell-Free Microtiter Plate Screen for Improved [FeFe]
Hydrogenases
James A. Stapleton
1
, James R. Swartz
1,2
*
1 Department of Chemical Engineering, Stanford University, Stanford, California, United States of America, 2 Department of Bioengineering, Stanford University, Stanford,
California, United States of America
Abstract
Background: [FeFe] hydrogenase enzymes catalyze the production and dissociation of H
2
, a potential renewable fuel.
Attempts to exploit these catalysts in engineered systems have been hindered by the biotechnologically inconvenient
properties of the natural enzymes, including their extreme oxygen sensitivity. Directed evolution has been used to improve
the characteristics of a range of natural catalysts, but has been largely unsuccessful for [FeFe] hydrogenases because of a
lack of convenient screening platforms.
Methodology/Principal Findings: Here we describe an in vitro screening technology for oxygen-tolerant and highly active
[FeFe] hydrogenases. Despite the complexity of the protocol, we demonstrate a level of reproducibility that allows
moderately improved mutants to be isolated. We have used the platform to identify a mutant of the Chlamydomonas
reinhardtii [FeFe] hydrogenase HydA1 with a specific activity ,4 times that of the wild-type enzyme.
Conclusions/Significance: Our results demonstrate the feasibility of using the screen presented here for large-scale efforts
to identify improved biocatalysts for energy applications. The system is based on our ability to activate these complex
enzymes in E. coli cell extracts, which allows unhindered access to the protein maturation and assay environment.
Citation: Stapleton JA, Swartz JR (2010) A Cell-Free Microtiter Plate Screen for Improved [FeFe] Hydrogenases. PLoS ONE 5(5): e10554. doi:10.1371/
journal.pone.0010554
Editor: Mark Isalan, Center for Genomic Regulation, Spain
Received March 7, 2010; Accepted April 9, 2010; Published May 10, 2010
Copyright:  2010 Stapleton, Swartz. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was funded by the Global Climate and Energy Project (GCEP) at Stanford University (http://gcep.stanford.edu/) as well as a National Defense
Science and Engineering (NDSEG) Fellowship (http://ndseg.asee.org/) to JAS. The funders had no role in study design, data collection and analysis, decision to
publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: jswartz@stanford.edu
Introduction
Hydrogenase enzymes reversibly catalyze the interconversion of
molecular hydrogen with protons and electrons [1]. They have
drawn increasing interest from the renewable energy community
[2,3] because of their potential to become key players in biological
hydrogen production schemes [4] and replacements for precious
metal catalysts in fuel cells [5,6]. Hydrogenases are classified as
[NiFe], [FeFe], or [Fe] according to the composition of their active
site metal cofactors. The [FeFe] hydrogenases have the highest
turnover rates [1], making them attractive for energy applications,
but are incompatible with many technologies due to their extreme
oxygen sensitivity [7,8]. Oxygen deactivates these hydrogenases by
diffusing into the core of the protein and reacting with the catalytic
iron-sulfur cofactor [9]. The development of an engineered [FeFe]
hydrogenase with improved industrial traits, including decreased
sensitivity to deactivation by oxygen and increased specific activity,
could make the economics of biological hydrogen production
technologies or fuel cells much more attractive.
Directed evolution, a laboratory technique that mimics natural
evolution by selecting the fittest mutants from large libraries [10],
is a promising method by which to improve the suitability of
hydrogenases for various biotechnological applications [11].
However, evolving hydrogenases has proved difficult due to a
lack of effective high-throughput screens. Developing a screen for
improved hydrogenase mutants is challenging because hydroge-
nases are difficult to express in active form and because their
substrate and product are difficult to measure in a high-
throughput format or to link to the survival of an organism. A
platform capable of supporting a large-scale screening effort has
yet to be described.
Heterologous expression of [FeFe] hydrogenases became
possible only recently, when the discovery of three helper proteins
required for synthesis and installation of the so-called H-cluster
cofactor at the catalytic site [12] enabled hydrogenase activation in
vivo in E. coli [13] and in vitro in E. coli cell extracts [14,15]. Prior to
this breakthrough, the low transformation efficiencies and slow
growth rates of the organisms that naturally harbor [FeFe]
hydrogenases prevented screening large in vitro-generated gene
libraries, and most directed evolution work focused on genome-
wide mutagenesis to improve hydrogen production rates or oxygen
tolerance of organisms that naturally produce hydrogen [16]. The
ability to express mutant [FeFe] hydrogenases in E. coli or by cell-
free protein synthesis (CFPS) has allowed the focus to shift to the
enzyme itself. Instead of evolving mutant organisms that might
shield their hydrogenases from oxygen or deliver reducing
equivalents to the protein more effectively, mutant hydrogenases
that are inherently oxygen tolerant and catalytically improved can
be developed. Such mutants would retain their improvements
independent of the choice of host organism, and analysis of their
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mutations could shed light on the relationship between [FeFe]
hydrogenase structure and function.
SIMPLEX (SIngle-Molecule PCR-Linked EXpression) [17] is
an entirely in vitro microtiter plate-based protein screening
platform. In vitro screens and selections, which use cell-free protein
synthesis to transform mutant DNA into mutant proteins, have a
number of advantages over in vivo methods. The protein synthesis
environment is open, allowing control over many aspects of the
protein production and folding environment that are inaccessible
in vivo. The protein product is synthesized within a few hours and
can be assayed without a cell lysis or permeablization step. In vitro
methods can be used to screen toxic proteins and can avoid many
of the biases for which cells are notorious. While much lower in
throughput than bulk methods such as ribosome display [18] and
in vitro compartmentalization [19], the microtiter plate format
allows a wide range of enzyme activities to be assessed using
standard colorimetric activity assays.
Here we present a SIMPLEX-based in vitro screen for [FeFe]
hydrogenase activity and oxygen tolerance (Figure 1), which we
have used to isolate a mutant of the Chlamydomonas reinhardtii [FeFe]
hydrogenase HydA1 with ,4-fold improved specific activity
relative to the wild-type. The system is conducive to automation
and can screen on the order of 10
3
–10
4
mutants per day.
A key step in SIMPLEX is single-molecule PCR (smPCR) [20],
which amplifies mutant genes isolated by limiting dilution. This
powerful technique is rapidly being adopted for use in a diverse
range of fields for applications such as cancer detection [21], gene
expression analysis [22], DNA methylation analysis [23], sequenc-
ing of unculturable strains [24], cell-free cloning [25], and directed
evolution. We present observations regarding factors that are
important for successful smPCR, which we hope will be of use to
researchers attempting to establish this powerful but delicate
technique in their own laboratories.
Results and Discussion
The screen for improved hydrogenase mutants consists of
mutant DNA library generation, limiting dilution, single-molecule
PCR, cell-free protein synthesis, oxygen exposure, spectrophoto-
metric measurement, and comparison to wild-type protein
(Figure 1).
Mutant DNA library generation
We chose the hydA1 gene encoding the HydA1 hydrogenase
from C. reinhardtii as our parent sequence, and generated a random
mutant DNA library using mutagenic PCR with nucleotide
analogs [26]. In general, libraries of linear mutant DNA templates
may be generated by any random or rational mutagenesis method,
including error-prone PCR [27], site-directed mutagenesis [28],
and recombination techniques [29,30]. The only requirements are
that the resulting mutant constructs contain homoprimer anneal-
ing sequences [31,32] to allow smPCR amplification [33] and an
appropriate promoter and ribosome binding site to allow
subsequent transcription and translation by the CFPS system
[34]. Sequencing of a sampling of mutant genes from the library
indicated a range of 2 to 10 DNA mutations per gene.
Limiting dilution and single-molecule PCR amplification
In the SIMPLEX system, each mutant gene is isolated from the
library by limiting dilution. Careful dilution to extremely low DNA
concentrations allows single molecules to be distributed into the
wells of a PCR plate according to Poisson distribution statistics.
We added mutant library DNA at an average of 2.3 molecules per
well, an amount that left approximately 10% of the wells empty,
which we felt represented an optimal compromise between
throughput and resolution. Decreasing the average number of
genes per well leaves more wells without DNA, while increasing
the average number of genes per well leads to signal distortion
when two or more genes are deposited and amplified in the same
well. The averaging of the signals from an improved mutant and
neutral or inactive mutants in the same well raises the threshold of
improvement required for detection. In addition, if the well is
found to contain improved enzymes, the genes must be separated
by limiting dilution or in vivo cloning before they can be retested
and sequenced.
The isolated genes are amplified to quantities sufficient for
CFPS reactions by single-molecule PCR. As the primer concen-
Figure 1. A schematic representation of the screening procedure.
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tration in smPCR is very high relative to that of the single template
molecule, primer-dimers and aberrant products are common. The
use of a homoprimer that anneals to both ends of the target
sequence has been shown to reduce primer dimerization [32] and
allow single-step single-molecule PCR [35]. Since any desired
primer sequence can be designed into the template, but not all
sequences are effective, we used the homoprimer sequences
reported by the developers of SIMPLEX (TR [36], K4 [37], and
SCA2 [38]), the efficacy of which we independently confirmed.
Accurately diluting DNA more than a billion fold to the single-
molecule level is not trivial. Since any template loss becomes very
significant at single-molecule concentrations, we performed the
dilutions in a buffered solution containing EDTA and blue dextran
[17] to block nonspecific adsorption of DNA. Solutions diluted to
a range of expected DNA concentrations were used as templates in
a series of PCRs until a dilution was found at which only a fraction
of the reactions amplified successfully. Additional amplifications
were then performed to more precisely estimate the DNA
concentration in that dilution by comparison of amplification
frequencies with the predictions of the Poisson distribution.
Though a variety of buffers and a wide range of concentrations
of magnesium, nucleotide triphosphates, and primers have been
recommended by reports in the literature, we were successful using
buffers supplied with commercial polymerases and Mg
2+
, dNTPs,
and primer concentrations typical of standard PCR protocols. We
also evaluated the addition of colloidal gold nanoparticles, which
have been reported to improve PCR specificity [32]. No
improvement was observed relative to reactions from which gold
was omitted. The volume of each smPCR reaction was 7 mL,
minimizing polymerase cost while ensuring that the liquid volume
would not be significantly depleted by evaporation during the long
thermocycling program. The melting step in our protocol was
shortened to 10 seconds at 95uC, and even shorter times are
sufficient for most targets. Minimizing the duration of this high-
temperature step minimized polymerase deactivation during the
high number of cycles required to amplify a single molecule to
saturation. The 80-cycle program was complete in about five
hours.
The sensitivity that allows amplification of a single copy of a
target also makes smPCR susceptible to contamination by even
very small amounts of DNA. While the specificity of the
homoprimer allows smPCR to tolerate contamination by envi-
ronmental or other nonspecific DNA, contamination by DNA
with homoprimer annealing sites can be a serious problem.
Because of this, extreme caution must be taken to avoid
contaminating reagents, pipettes, surfaces, and consumables with
template or product DNA. To avoid contamination, all smPCR
reaction solutions were mixed in one room, and the products
analyzed in a second room. Reaction components were mixed in a
laminar flow hood with a dedicated set of pipettes using smPCR-
only reagent aliquots, which were frequently changed. Despite
these precautions, contamination was a frequent occurrence, and
negative controls without template were routinely run to ensure
that the reactions were contaminant-free.
Comparison of observed results with the predictions of the
Poisson distribution confirmed that amplified products originated
from single template molecules and allowed determination of the
number of DNA molecules in the stock solution (Table 1). We
prepared a mixture consisting of PCR reagents and a known
volume of a highly diluted template solution. We then divided a
96-well PCR plate into thirds, and added increasing volumes of
the mixture (5, 10, and 15 mL) into the wells of each section,
reserving three wells for no-template controls. Within each section
of the plate, observation of the fraction of the reactions in which
amplification occurred allowed us to estimate the number of DNA
molecules in the volume of template solution in the reaction
mixture. Agreement across the sections indicated successful
smPCR.
Cell-free protein synthesis
Following amplification of isolated mutant DNA templates,
smPCR plates were brought into an anaerobic chamber, and
1.5 mL volumes (containing approximately 100 ng [34,36] of
DNA) were transferred into the corresponding wells of a CFPS
reaction plate without purification. The PCR plates, containing
the remainder of each PCR product, were then archived at
220uC so that DNA corresponding to eventual ‘‘hits’’ interesting
enough to merit further evaluation could be easily recovered.
Extensive modifications have been made to the PANOx-SP [39]
CFPS protocol developed for general use in our laboratory to
allow activation of [FeFe] hydrogenases [14]. We prepared the
extracts from E. coli cells in which the three maturase proteins
required for synthesis and activation of the [FeFe] hydrogenase
active site [12] had been heterologously expressed. We conducted
the CFPS reactions anaerobically to prevent deactivation of
hydrogenase and its maturases. To stabilize the linear templates
produced by PCR-based mutagenesis techniques, we added
purified lambda phage Gam protein to the reaction mixtures to
inhibit the RecBCD exonuclease complex [40] present in the
extract. We also supplemented the reactions with S-adenosylme-
thionine (SAM), a substrate of the maturases [41]. The reactions
are capable of synthesizing and maturing 40 ng/mL of active
hydrogenase.
Multiplexed hydrogenase activity assays
Methyl viologen is a redox-sensitive dye that is clear when
oxidized and blue when reduced, and which can exchange
electrons with [FeFe] hydrogenases. Spectrophotometric measure-
ment of the rate of the color change as hydrogenase oxidizes H
2
and reduces methyl viologen has been used widely in the literature
as an assay for hydrogenase activity [42], and we adopted it as our
primary activity assay. Preliminary hits identified with this screen
could be assayed with the natural electron donor, the protein
ferredoxin, in a hydrogen production assay to ensure compatibility
before being carried on to the next round of mutagenesis.
We mixed the CFPS reaction products with a buffered methyl
viologen solution. The absorbance at 578 nm in each well of a 96-
well microtiter plate was measured over two minutes with a plate
reader within the anaerobic chamber as hydrogenase consumed
dissolved H
2
. The slope of the absorbance change with time was
converted into a hydrogenase activity measurement using Beer’s
Law. The absorbance of methyl viologen solutions in this H
2
consumption assay remained constant in the absence of added
CFPS product, and the addition of negative control CFPS
mixtures incubated without any DNA template or with templates
Table 1. Comparison of experimental results with the Poisson
distribution.
Reaction size
Percent
amplified
Poisson average
molecules/reaction
Poisson average
molecules/5 mL
5 mL 26.4 0.31 0.31
10 mL 50 0.69 0.35
15 mL 62.3 0.98 0.33
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encoding chloramphenicol acetyltransferase (CAT) generated
slopes that were small compared to those generated by the
products of CFPS reactions expressing hydrogenase.
Multiplexed measurement of oxygen tolerance
To measure oxygen tolerance, we challenged the mutant
proteins by adding a liquid solution containing dissolved oxygen.
During the subsequent incubation, oxygen continually diffused out
of the liquid into the anaerobic atmosphere, and was also
consumed by the extract, presumably by oxygen-scavenging
cytochrome oxidases in inverted membrane vesicles [43]. The
oxygen concentration profile experienced by the hydrogenase
mutants was therefore not constant with time and could not be
quantitatively determined, but we presumed it to be the same for
each reaction mixture in a given microtiter plate. For the purpose
of high-throughput screening this oxygen exposure procedure
proved convenient and reliable. Loss into the atmosphere and
consumption by the extract eventually completely removed the
dissolved oxygen, which would otherwise have interfered with
measurement of the absorbance of the assay solution by oxidizing
the methyl viologen.
The extent of the deactivation could be controlled by adjusting
the volume of oxygen-containing liquid added (Figure 2). Weak
deactivation resulted in a higher signal-to-noise ratio, but led to
high sensitivity to unintended variations in exposure. After very
strong deactivation, the residual activity was obscured by
background absorbance changes caused by factors within the cell
extract. A post-exposure residual activity of 15–20% avoided both
problems, generating a signal several times greater than the
background observed in CAT control reactions.
The volume of CFPS reaction product used for the post-
exposure activity measurement was adjusted such that the slopes of
the pre- and post-exposure measurements were roughly equal and
within the range that allowed accurate activity determination. For
example, in the case of a 20% residual activity target, five times as
much reaction product was used in the post-exposure assay as in
the pre-exposure assay.
Data processing and hit identification
The oxygen tolerance screen included two activity measure-
ments of each CFPS reaction product with the methyl viologen
activity assay, one before oxygen exposure and another after. The
ratio of the two measurements was used as a measure of oxygen
tolerance and was the criterion by which hits were identified. The
pre-exposure test allowed the oxygen tolerance score of each
mutant to be normalized by initial activity, eliminating the
influence of variation in expression across CFPS reactions. In
addition, a minimum pre-exposure activity cutoff was established.
Mutant hydrogenases with pre-exposure activities below this
threshold were disqualified, since the low signal-to-noise ratio in
these wells inflated the residual activity ratio.
The activity and residual activity ratio of each mutant were
compared to those of wild-type hydrogenase controls. Performing
wild-type control reactions in each microtiter plate enabled us to
account for plate-to-plate and day-to-day variations in CFPS
reaction performance and oxygen exposure effectiveness. The
ability of a screen to identify an improved mutant is inversely
related to the standard deviation of measurements of identically
prepared wild-type reactions. We were able achieve a coefficient of
variance of measurements of oxygen tolerance of ,15–25%. The
precision of repeated measurements of oxygen tolerance is shown
in Figure 3. Experiments 1 and 2 measured the error of residual
activity measurements on the products of identically performed
CFPS reactions expressing wild-type C. reinhardtii HydA1. In
Experiment 3 wild-type C. pasteurianum hydA was used as the
Figure 2. Oxygen deactivation curve for the product of CFPS
reactions expressing C. reinhardtii HydA1. 5 mL of CFPS product
were diluted with 25 mL of anaerobic Tris buffer before addition of the
volume of air-saturated Tris buffer indicated on the x-axis. The slope of
methyl viologen absorbance at 578 nm with time for each oxygen-
exposed sample was normalized to the slope with no oxygen exposure.
The pink curve represents an exponential decay fit to the data. Error
bars indicate the standard deviation of triplicate measurements.
doi:10.1371/journal.pone.0010554.g002
Figure 3. Assessment of the well-to-well error of the screen in
three sets of identically performed 10 mL CFPS reactions.
Residual activity following oxygen exposure is represented by the
average pre-exposure/post-exposure activity ratio. Error bars indicate
the standard deviation for n = 88, 72, and 71 wells respectively. The
coefficient of variance is given above each bar. 1 and 5 mL of CFPS
product were assayed in the pre-exposure and post-exposure
measurements, respectively, and the post-exposure activity was
normalized to 1 mL. 15 mLofO
2
-equilibrated buffer were used for
deactivation. C. reinhardtii HydA1 was expressed in Experiments 1 and
2, and C. pasteurianum CpI was expressed in Experiment 3. The
difference in average residual activity between Experiments 1 and 2 is
likely due to differences in the extent of oxygen diffusion out of the
aerobic buffer before addition to the hydrogenase solutions, and
highlights the need for wild-type controls on each plate.
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template. The measured error is in a range typical of microtiter
plate screens. If a hit threshold were set two standard deviations
above the mean, the screen could identify mutants improved in
oxygen tolerance by ,30–50%.
Isolation and characterization of a mutant with improved
specific activity
We used this method to screen ,30,000 hydA1 mutants (an
average of 2 mutants per well of 150 96-well plates). We found no
hydrogenase mutants that were significantly more oxygen-tolerant
than the wild-type. However, pre-exposure hydrogen consumption
activity measurements of several mutants were significantly higher
than wild-type HydA1 activity measurements, and these mutants
were selected for further evaluation. The DNA templates were
recovered from archived PCR plates, re-amplified by PCR, and
evaluated by expression in plate-based CFPS followed by activity
quantification of the protein products with the methyl viologen
activity assay.
To measure the specific activity of the most improved mutant,
we added the radioactive amino acid L-[U–
14
C]-leucine to the
CFPS mixtures, and measured its incorporation into the protein
product by TCA precipitation and liquid scintillation counting
[44]. We intentionally limited the protein yields in these reactions
to promote complete activation, and their proximity to the
sensitivity limit of our protein quantification technique led to a
significant standard deviation among the measurements. We
calculated the specific activity in the hydrogen production
direction after measuring hydrogen produced over time by CFPS
product mixed with sodium dithionite-reduced methyl viologen
using an H
2
analyzer [14].
The most improved HydA1 mutant is more active than the
wild-type by a factor of ,4 (Figure 4). Interestingly, the activity of
the mutant seems to have increased by the same amount in both
directions, indicating that its mutations do not bias its reversibility
[45]. The specific activity of the wild-type HydA1 is 150 pmol H
2
/
min/ng protein. This value is ,20% of typical activities reported
for HydA1 purified after production in hosts naturally possessing
[FeFe] hydrogenases [46–49], but very similar to specific activities
reported for protein produced in vivo in E. coli [13]. Because of this
discrepancy, we cannot rule out the possibility that the isolated
mutant is improved in its ability to be activated in the CFPS
system rather than in its turnover rate. To confirm the
improvement, future work will include expression of the mutant
in a natural [FeFe] hydrogenase host in place of the wild-type
gene.
DNA sequencing revealed two amino acid mutations: G172D
and N267S (residue numbers include the 55 amino acid targeting
sequence, as in GenBank accession AY055755.1). A multiple
sequence alignment generated by ClustalW (Figure S1) showed
that both of these amino acids are immediately adjacent in the
primary sequence to residues that are highly conserved among
[FeFe] hydrogenases. Interestingly, both are non-conservative
changes. No crystal structure is available for C. reinhardtii HydA1,
but the position corresponding to G172 in the homologous
hydrogenase CpI from Clostridium pasteurianum [50] is in an alpha
helix approximately 8A
˚
from the active site H-cluster, while the
residue corresponding to N267 is on an external loop.
This is the first improved [FeFe]hydrogenasetobeisolated
by directed evolution. Previously, a mutant of the large subunit
of the E. coli [NiFe] hydrogenase 3 was found that improved the
hydrogen production capability of its host cell [51]. This
membrane-bound hydrogenase could not be assayed in purified
form. The mutant was identified with an in vivo screen using
chemochromic hydrogen detectors, with a throughput theoret-
ically comparable to that of the screen presented here. Recent
progress in the in vivo expression of [FeFe] hydrogenases in E.
coli couldallowthisorganismtobeusedasahostforin vivo
screening of heterologous [FeFe] hydrogenases using a similar
system.
Although we developed the screen to search for oxygen-
tolerant mutants, it can also be used to identify mutants with
increased specific activity, as well as mutants improved in any
other activity-related trait. For example, thermostable mutants
could be identified by incubating the CFPS products at high
temperatures and then measuring residual activity. Since the
screen is based on cell-free protein synthesis of [FeFe]
hydrogenase mutants, the experimenter has unfettered access to
the protein synthesis, maturation, and activity assay environment,
providing flexibility in devising screening conditions that would
not be possible in traditional in vivo platforms. The cell-free nature
of the screen may prove especially advantageous when screening
for characteristics beneficial for in vitro applications such as fuel
cells.
Our results further validate SIMPLEX as a valuable technique
in the repertoire of directed evolution methods. The HydA1
hydrogenase from C. reinhardtii is a difficult target for both smPCR,
because of the 60% GC content of the gene, and CFPS, because of
the complexity of the protein. The success of SIMPLEX in
evolving this protein validates the system for use evolving other
complex, industrially-relevant enzymes.
The ability of our screen to identify mutants with improved
specific activities, while simultaneously failing to identify mutants
with improved oxygen tolerance, hints at the relative frequency of
the two phenotypes in the mutant library. It is possible that any
improvement in oxygen tolerance will require multiple simulta-
neous mutations to close off all of the routes by which oxygen can
reach the active cluster. If this is the case, either new screens with
higher throughputs or a massive effort with a screen such as the
one presented here will be required, along with rational or semi-
rational library design [52] to restrict the sequence space to be
searched.
Figure 4. Specific activities of wild-type and mutant C.
reinhardtii HydA1. Specific activity was calculated for each using
cell-free protein synthesis products in the hydrogen consumption
direction (blue bars) and the hydrogen production direction (red bars).
Error bars indicate the standard deviation of at least n = 4 independent
experiments.
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Materials and Methods
Template preparation, purification, and dilution
DNA templates for smPCR were linear cell-free protein
synthesis expression templates consisting of a codon-optimized
gene encoding C. reinhardtii HydA1 hydrogenase with a T7 RNA
polymerase promoter and terminator and an E. coli ribosome
binding site [14]. The 55 amino-acid targeting sequence following
the N-terminal methionine was removed, and the first nine codons
of the chloramphenicol acetyltransferase gene, which has been
shown to express well in the CFPS system, were inserted after the
start codon to ensure accessibility of the RBS on the mRNA
transcript. The templates were amplified from a plasmid using
primers that annealed to sequences on the plasmid outside the
expression region, and included 59 regions that extended the
homoprimer annealing sequence of choice onto each end of the
linear product. Nucleotide analog mutagenesis [26] was used to
generate mutant libraries.
Templates were purified with QiaQuick PCR cleanup kits
(Qiagen, Valencia, CA), by purification from Novex TBE
polyacrylamide gels (Invitrogen, Carlsbad, CA), or by phenyl
chloroform extraction and ethanol precipitation.
DNA was quantified by measuring absorption at 260 nm with a
spectrophotometer. Desired dilution levels were calculated based
on the calculated molecular weight of the template molecules
(based on an average of 660 Da/bp). DNA was serially diluted
into TE buffer with 0.1% (w/v) blue dextran, usually in steps of
1 mL into 1 mL. Tubes were thoroughly vortexed between dilution
steps. DNA stocks were diluted to an estimated concentration of
100 molecules/mL for use in SIMPLEX screens. The concentra-
tions of working stocks were confirmed by Poisson distribution
experiments as described, and the amount added to the smPCR
mixture was adjusted as necessary to achieve the desired number
of molecules per well of a PCR plate.
Single-molecule PCR
Single-molecule PCR reaction mixtures consisted of 1X
manufacturer’s PCR buffer (supplemented with magnesium
chloride to 2 mM when necessary); 0.2 mM of each of the four
deoxyribonucleotides; 0.5 mM of the homoprimer; either 0.02
units/mL Platinum Taq (Stratagene, La Jolla, CA), 0.03 units/mL
Pfu Turbo (Stratagene, La Jolla, CA), 0.03 units/mL Pfu Ultra
(Stratagene, La Jolla, CA), or 0.02 units/mL Phusion (Finnzymes,
Espoo, Finland); and template DNA. Reaction volumes were
typically 7 mL. Reactions were prepared in a laminar flow hood
(Nuaire, Plymouth, MN) to prevent contamination.
Reactions were incubated at 95uC for one minute to activate the
hot-start polymerase and denature the template, then thermo-
cycled 80 times through 10 seconds at 95uC, 20 seconds at 55uC,
and an extension temperature and time recommended by the
manufacturer (generally 72uC for 1 minute/kb of template length).
When Phusion polymerase was used, the denaturation step was 5
seconds at 98uC, the annealing temperature was 62uC, and the
extension time was 15 seconds/kilobase.
Cell-free protein synthesis
Extracts were prepared anaerobically from E. coli BL21 cultures
expressing the three helper proteins required for maturation of
[FeFe] hydrogenases [14]. Cells were grown aerobically on a
defined medium [14] at 37uC in a fermenter to OD ,6–8. The
culture was then made anaerobic by switching the gas feed from
air to argon. After 45 minutes, the temperature was dropped to
15uC, and expression of the helper proteins was induced by
addition of 0.5 mM IPTG. At this point the medium was also
supplemented with 0.33 mg/mL ferric ammonium citrate and
10 mM fumarate. After 16 hours the cells were harvested,
pelleted, washed, and homogenized, and the lysate centrifuged
and frozen, as described previously [14] except that all the steps
were performed under aerobic conditions. Before use in CFPS
reactions, the extract was incubated at room temperature with
1 mM ferrous ammonium sulfate and 1 mM sodium sulfide for 1–
2 hours [14].
CFPS reactions were conducted in 384-well PCR plates. 7.5 mL
reaction mixtures were prepared by adding 3 mL of Mix 1 to a
well, then 1.5 mL of crude smPCR product, then 3 mL of Mix 2.
Mix 1 consisted of small molecule cofactors (listed in Table S1 at
their concentrations in the final CFPS mixture). Mix 2 consisted of
the following, given with their concentrations in the final CFPS
mixture: 0.1 mg/mL T7 RNA polymerase (overexpressed in E. coli
and purified in house), 6.7 mg/mL lambda phage Gam protein
(expressed in CFPS and purified in house), 0.01% 31R1 antifoam
(Sigma Aldrich, St Louis, MO), and 25% v/v cell extract.
14
C-
leucine was omitted in the high-throughput screen reactions but
included at 5.25 mM when characterizing hits to allow quantifi-
cation of synthesized protein by scintillation counting of
incorporated radiation [44]. Reactions were incubated for
between six and 16 hours at room temperature in an anaerobic
glove box (Coy Laboratory Products, Grass Lake, MI) before
being assayed.
Methyl viologen activity assay
Activity of the hydrogenase produced in the CFPS reactions was
obtained by measuring the rate of change of the absorbance at
578 nm of 200 mL of a 2 mM solution of methyl viologen in
50 mM Tris buffer at pH 8 with a VersaMax plate reader
(Molecular Devices, Sunnyvale, CA) following addition of CFPS
product. Immediately before the assay was conducted, the methyl
viologen solution was reduced by addition of titanium citrate until
a light blue color persisted. Absorbance was monitored for two
minutes at room temperature in 96-well flat-bottom plates. Slopes
were converted into activity using Beer’s Law with an extinction
coefficient of 9.78 AU/min/mM.
Oxygen exposure
50 mM Tris buffer at pH 8 was stored outside the glove box
and allowed to come to equilibrium with oxygen, so that the
dissolved oxygen concentration was expected to be ,0.25 mM. A
small vial was filled to the brim with this solution to minimize
headspace, sealed, and moved into the anaerobic chamber
immediately before the oxygen exposure step in the protocol.
The vial was opened and the buffer poured into a trough, from
which it was aspirated by a multichannel pipetter or the
multichannel head of a liquid handler. A volume of buffer
expected to deactivate the CFPS product to the desired extent was
dispensed into each well of the oxygen-exposure plate. The plate
was incubated for 10 minutes, after which time methyl viologen
could be added to assay residual activity. No decrease in blue color
was noticeable upon methyl viologen addition, indicating that after
10 minutes all the oxygen had either been consumed by the extract
or had diffused into the atmosphere of the anaerobic chamber.
High-throughput activity measurements
A liquid handling machine (EpMotion 5070, Eppendorf North
America, Westbury, NY) first pipetted 72 mL of anaerobic Tris
buffer (50 mM, pH 8) into each well of eight 96-well spectropho-
tometer plates, four for pre-oxygen exposure and four for post-
oxygen exposure activity measurements of the mutant proteins in
one 384-well CFPS plate. 6 mL of CFPS product were transferred
A Cell-Free Hydrogenase Screen
PLoS ONE | www.plosone.org 6 May 2010 | Volume 5 | Issue 5 | e10554

from the 384-well CFPS reaction plate into the wells of four of the
96-well plates, and mixed with the buffer by pipetting up and
down five times. 13 mL were then transferred from these wells to
the corresponding wells of the other four plates, and again mixed.
These plates, which would be used for the pre-oxygen exposure
measurements, thus contained 1/5 as much CFPS product as did
the first set of plates, which would be used for the post-oxygen
exposure measurements. After the liquid handler dispensed 20 mL
of air-equilibrated buffer into each well of the post-deactivation
plates, the wells of all eight plates contained 85 mL of liquid.
115 mL of methyl viologen solution was then added to each well to
give a final concentration of 2 mM, and the plate was immediately
inserted into the spectrophotometer.
Hit confirmation and characterization
Genes were reamplified by PCR and re-tested in replicate CFPS
reactions. Radioactive leucine was added to allow quantification of
protein production by TCA precipitation and liquid scintillation
counting [44]. Specific activity in the hydrogen consumption
direction was calculated by dividing the hydrogen consumption
rate determined from the methyl viologen assay data by the
soluble protein yield determined by measurement of incorporated
radioactive leucine. Mutants were tested for hydrogen production
ability using a Peak Performer hydrogen analyzer (Peak Labora-
tories, Mountain View, CA) as previously reported [14]. Crude
CFPS solution containing 10–40 ng of hydrogenase was added to
1 mL of a solution consisting of 5 mM methyl viologen and
25 mM sodium dithionite in 50 mM Tris-HCl pH 6.8 in a sealed
vial. The headspace was exchanged with nitrogen, and 100 mL
samples were removed from the headspace with a syringe and
injected into the hydrogen analyzer. Specific activity in the
hydrogen production direction was calculated by dividing the
hydrogen production rate calculated from the hydrogen analyzer
data by the soluble protein yield calculated by measurement of
incorporated radioactive leucine. Due to the observed dependence
of the percentage of hydrogenase that is matured on the expression
level, specific activities were only compared when the expression
levels of each sample were similar.
For molecular modeling, we used the PyMOL Molecular
Graphics System with PDB entry 1feh.
Supporting Information
Figure S1 Multiple sequence alignment of representative [FeFe]
hydrogenases. Asterisks, colons, and periods indicate locations
with homology across different species. Red dots indicate the
locations of the mutations identified in this study. Both mutations
are adjacent to highly conserved residues. CpI: HydA from
Clostridium pasteurianum. C.r.HydA1 and C.r.HydA2: HydA1 and
HydA2 from Chlamydomonas reinhardtii.
Found at: doi:10.1371/journal.pone.0010554.s001 (2.64 MB TIF)
Table S1 Final concentrations of small molecule components of
the CFPS mixture.
Found at: doi:10.1371/journal.pone.0010554.s002 (0.04 MB
DOC)
Acknowledgments
The authors thank Sean Kendall for his assistance performing the screen
and Jon Kuchenreuther and Bertrand Lui for comments on the
manuscript.
Author Contributions
Conceived and designed the experiments: JAS JRS. Performed the
experiments: JAS. Analyzed the data: JAS JRS. Contributed reagents/
materials/analysis tools: JRS. Wrote the paper: JAS.
References
1. Adams MW (1990) The structure and mechanism of iron-hydrogenases.
Biochimica et Biophysica Acta 1020: 115–145.
2. Mertens R, Liese A (2004) Biotechnological applications of hydrogenases.
Current Opinion in Biotechnology 15: 343–348.
3. Levin D, Pitt L, Love M (2004) Biohydrogen production: prospects and
limitations to practical application. International Journal of Hydrogen Energy:
29: 173 –185.
4. Prince RC, Kheshgi HS (2005) The photobiological production of hydrogen:
potential efficiency and effectiveness as a renewable fuel. Critical Reviews in
Microbiology 31: 19–31.
5. Tye JW, Hall MB, Darensbourg MY (2005) Better than platinum? Fuel cells
energized by enzymes. Proc Natl Acad Sci U S A 102: 16911–16912.
6. Vincent KA, Parkin A, Armstrong FA (2007) Investigating and exploiting the
electrocatalytic properties of hydrogenases. Chemical Reviews 107: 4366–4413.
7. Abeles FB (1964) Cell-free hydrogenase from Chlamydomonas. Plant Physiology
39: 169–176.
8. Erbes DL, King D, Gibbs M (1979) Inactivation of hydrogenase in cell-free
extracts and whole cells of Chlamydomonas reinhardtii by oxygen. Plant Physiology
63: 1138–1142.
9. Cohen J, Kim K, Posewitz M, Ghirardi ML, Schulten K, et al. (2005) Molecular
dynamics and experimental investigation of H
2
and O
2
diffusion in [Fe]-
hydrogenase. Biochem Soc Trans 33: 80–82.
10. Arnold FH (1998) Design by directed evolution. Accounts of Chemical Research
31: 125–131.
11. Turner NJ (2009) Directed evolution drives the next generation of biocatalysts.
Nature Chemical Biology 5: 567–73.
12. Posewitz MC, King PW, Smolinski SL, Zhang L, Seibert M, et al. (2004)
Discovery of two novel radical S-adenosylmethionine proteins required for the
assembly of an active [Fe] hydrogenase. The Journal of Biological Chemistry
279: 25711–20.
13. King P, Posewitz M, Ghirardi M, Seibert M (2006) Functional studies of [FeFe]
hydrogenase maturation in an Escherichia coli biosynthetic system. Journal of
Bacteriology 188: 2163–2172.
14. Boyer M, Stapleton JA, Kuchenreuther JM, Wang C, Swartz JR (2008) Cell-free
synthesis and maturation of hydrogenases. Biotechnology and Bioengineering
99: 59–67.
15. Kuchenreuther JM, Stapleton JA, Swartz JR (2009) Tyrosine, cysteine, and S-
adenosyl methionine stimulate in vitro [FeFe] hydrogenase activation. PloS ONE
4(10): e7565. DOI:10.1371/journal.pone.0007565.
16. Flynn T, Ghirardi ML, Seibert M (2002) Accumulation of O
2
-tolerant
phenotypes in H
2
-producing strains of Chlamydomonas reinhardtii by sequential
applications of chemical mutagenesis and selection. International Journal of
Hydrogen Energy 27: 1421–1430.
17. Rungpragayphan S, Nakano H, Yamane T (2003) PCR-linked in vitro
expression: a novel system for high-throughput construction and screening of
protein libraries. FEBS Letters 540: 147–150.
18. Hanes J, Pluckthun A (1997) In vitro selection and evolution of functional
proteins by using ribosome display. Proc Natl Acad Sci U S A 94: 4937–
4942.
19. Tawfik DS, Griffiths AD (1998) Man-made cell-like compartments for molecular
evolution. Nature Biotechnology 16: 652–656.
20. McCaughan F, Dear PH (2009) Single-molecule genomics. The Journal of
Pathology DOI:10.1002/path.2647.
21. Vogelstein B, Kinzler KW (1999) Digital PCR. Proc Natl Acad Sci U S A 96:
9236–9241.
22. Pohl G, Shih I (2004) Principle and applications of digital PCR. Expert Review
of Molecular Diagnostics 4: 41–47.
23. Chhibber A, Schroeder BG (2008) Single-molecule polymerase chain reaction
reduces bias: application to DNA methylation analysis by bisulfite sequencing.
Analytical Biochemistry 377: 46–54.
24. Hutchison III CA, Venter JC (2006) Single-cell genomics. Nature Biotechnology
24: 657–658.
25. Kraytsberg Y, Khrapko K (2005) Single-molecule PCR: an artifact-free PCR
approach for the analysis of somatic mutations. Expert Review of Molecular
Diagnostics 5: 809–815.
26. Zaccolo M, Williams DM, Brown DM, Gherardi E (1996) An approach to
random mutagenesis of DNA using mixtures of triphosphate derivatives of
nucleoside analogues. Journal of Molecular Biology 255: 589–603.
27. Cirino PC, Mayer KM, Umeno D (2003) Generating mutant libraries using
error-prone PCR. In: Arnold FH, Georgiou G, eds. Methods in Molecular
Biology, vol 231: Directed Evolution Library Creation: Methods and Protocols.
Totowa, NJ: Humana Press. pp 3–9.
A Cell-Free Hydrogenase Screen
PLoS ONE | www.plosone.org 7 May 2010 | Volume 5 | Issue 5 | e10554

28. Ho SN, Hunt HD, Horton RM, Pullen JK, Pease LR (1989) Site-directed
mutagenesis by overlap extension using the polymerase chain reaction. Gene 77:
51–9.
29. Crameri A, Raillard SA, Bermudez E, Stemmer WP (1998) DNA shuffling of a
family of genes from diverse species accelerates directed evolution. Nature 391:
288–91.
30. Nagy LE, Meuser LE, Plummer S, Seibert M, Ghirardi ML, et al. (2007)
Application of gene-shuffling for the rapid generation of novel [FeFe]-
hydrogenase libraries. Biotechnol Lett DOI: 10.1007/s10529-006-9254-9.
31. Rungpragayphan S, Yamane T, Nakano H (2007) SIMPLEX: single-molecule
PCR-linked in vitro expression: a novel method for high-throughput construction
and screening of protein libraries. In: Grandi G, ed. Methods in Molecular
Biology, vol 375: In Vitro Transcription and Translation Protocols, Second
Edition. Totowa, NJ: Humana Press. pp 79–94.
32. Brownie J, Shawcross S, Theaker J, Whitcombe D, Ferrie R, et al. (1997) The
elimination of primer-dimer accumulation in PCR. Nucleic Acids Research 25:
3235–3241.
33. Ohuchi S, Nakano H, Yamane T (1998) In vitro method for the generation of
protein libraries using PCR amplification of a single DNA molecule and coupled
transcription/translation. Nucleic Acids Research 26: 4339–4346.
34. Woodrow KA, Airen IO, Swartz JR (2006) Rapid expression of functional
genomic libraries. Journal of Proteome Research 5: 3288–3300.
35. Nakano H, Kobayashi K, Ohuchi S, Sekiguchi S, Yamane T (2000) Single-step
single-molecule PCR of DNA with a homo-priming sequence using a single
primer and hot-startable DNA polymerase. Journal of Bioscience and
Bioengineering 90: 456–458.
36. Rungpragayphan S, Kawarasaki Y, Imaeda T, Kohda K, Nakano H, et al.
(2002) High-throughput, cloning-independent protein library construction by
combining single-molecule DNA amplification with in vitro expression. Journal of
Molecular Biology 318: 395–405.
37. Koga Y, Yamane T, Nakano H (2007) Creation of novel enantioselective lipases
by SIMPLEX. In: Grandi G, ed. Methods in Molecular Biology, vol 375: In Vitro
Transcription and Translation Protocols, Second Edition. Totowa, NJ: Humana
Press. pp 165–181.
38. Rungpragayphan S, Haba M, Nakano H, Yamane T (2004) Rapid screening for
affinity-improved scFvs by means of single-molecule-PCR-linked in vitro
expression. Journal of Molecular Catalysis B: Enzymatic 28: 223–228.
39. Jewett MC, Swartz JR (2004) Mimicking the Escherichia coli cytoplasmic
environment activates long-lived and efficient cell-free protein synthesis.
Biotechnology and Bioengineering 86: 19–26.
40. Sitaraman K, Esposito D, Klarmann G, Le Grice SF, Hartley JL, et al. (2004) A
novel cell-free protein synthesis system. Journal of Biotechnology 110: 257–263.
41. Rubach JK, Brazzolotto X, Gaillard J, Fontecave M (2005) Biochemical
characterization of the HydE and HydG iron-only hydrogenase maturation
enzymes from Thermatoga maritima. FEBS letters 579: 5055–5060.
42. Peck H, Gest H (1956) A new procedure for assay of bacterial hydrogenases.
Journal of Bacteriology 71: 70–80.
43. Jewett MC, Calhoun KA, Voloshin A, Wuu JJ, Swartz JR (2008) An integrated
cell-free metabolic platform for protein production and synthetic biology.
Molecular Systems Biology 4: 220.
44. Kim D, Swartz J (2001) Regeneration of adenosine triphosphate from glycolytic
intermediates for cell-free protein. Biotechnology and Bioengineering 74:
309–316.
45. McTavish H, Sayavedra-Soto LA, Arp DJ (1995) Substitution of Azobacter
vinelandii hydrogenase small-subunit cysteines by serines can create insensitivity
to inhibition by O
2
and preferentially damages H
2
oxidation over H
2
evolution.
Journal of Bacteriology 177: 3960–3964.
46. Roessler P, Lien S (1984) Purification of hydrogenase from Chlamydomonas
reinhardtii. Plant Physiology 75: 705–709.
47. Sybirna K, Antoine T, Lindberg P, Fourmond V, Rousset M, et al. (2008)
Shewanella oneidensis: a new and efficient system for expression and maturation of
heterologous [Fe-Fe] hydrogenase from Chlamydomonas reinhardtii. BMC Biotech-
nology 8: 73.
48. Florin L, Tsokoglou A, Happe T (2001) A novel type of iron hydrogenase in the
green alga Scenedesmus obliquus is linked to the photosynthetic electron transport
chain. Biochemistry 276: 6125–6132.
49. Happe T, Naber J (1993) Isolation, characterization and N-terminal amino acid
sequence of hydrogenase from the green alga Chlamydomonas reinhardtii. FEBS
Journal 214: 475.
50. Peters JW, Lanzilotta WN, Lemon BJ, Seefeldt LC (1998) X-ray crystal structure
of the Fe-only hydrogenase (CpI) from Clostridium pasteurianum to 1.8 angstrom
resolution. Science 282: 1853–1858.
51. Maeda T, Sanchez-Torres V, Wood TK (2008) Protein engineering of
hydrogenase 3 to enhance hydrogen production. Applied Microbiology and
Biotechnology 79: 77–86.
52. Liebgott P, Leroux F, Burlat B, Dementin S, Baffert C, et al. (2010) Relating
diffusion along the substrate tunnel and oxygen sensitivity in hydrogenase.
Nature Chemical Biology 6: 63–70.
A Cell-Free Hydrogenase Screen
PLoS ONE | www.plosone.org 8 May 2010 | Volume 5 | Issue 5 | e10554