ARTICLE
Cell-Free Synthesis and Maturation
of [FeFe] Hydrogenases
1 1 eut
38
55;
am
OI 1
2002), cytochrome P450’s (Vail et al., 2005), and hydro-
two electrons to form molecular hydrogen. Relative to the
possible alternative energy
nique iron-sulfur cluster, the
requires CO and CN ligands
ed to be di(thiomethyl)amine
conserved set of maturation
ydG (in eukaryotes HydE
ingle protein, HydEF), have
for the activation of [FeFe]
Biotechnology and Bioengineering, Vol. 99, No. 1, January 1, 2008 59This article contains Supplementary Material available at
http://www.interscience.wiley.com/jpages/0006-3592/suppmat.
Correspondence to: J.R. Swartz

2007 Wiley Periodicals, Inc.providing control over the maturation conditions. These
new capabilities will enhance the investigation of complex
proteins requiring helper proteins for maturation and move
us closer to the development of improved hydrogenases for
biological production of hydrogen as a clean, renewable
alternative fuel.
Biotechnol. Bioeng. 2008;99: 59–67.

2007 Wiley Periodicals, Inc.
genases have very fast hydroge
attracted much attention for
applications. In addition to a u
active center of these proteins
and a bridging molecule propos
(Fan and Hall, 2001). A highly
enzymes, HydE, HydF, and H
and HydF are fused into a s
been shown to be necessaryof cell walls permits direct addition of cofactors and sub-
strates, enabling rapid production of active protein and phylogenetically distinct [NiFe] hydrogenases, [FeFe] hydro-
n production rates and haveMarcus E. Boyer, James A. Stapleton, Jon M. Kuchenr
Chia-wei Wang,1 James R. Swartz1,2
1Department of Chemical Engineering, Stanford University,
California 94305; telephone: 650-723-5398; fax: 650-725-05
2Department of Bioengineering, Stanford University, 318 C
California 94305
Received 23 December 2006; revision received 11 May 2007; accepted 14 May 2007
Published online 1 June 2007 in Wiley InterScience (www.interscience.wiley.com). D
ABSTRACT: [FeFe] hydrogenases catalyze the reversible
reduction of protons to molecular hydrogen (Adams
(1990); Biochim Biophys Acta 1020(2): 115–145) and are
of significant interest for the biological production of
hydrogen fuel. They are complex proteins with active sites
containing iron, sulfur, and carbon monoxide and cyanide
ligands (Peters et al. (1998); Science 282(5395): 1853–1858).
Maturation enzymes for [FeFe] hydrogenases have been
identified (Posewitz et al. (2004); J Biol Chem 279(24):
25711–25720), but complete mechanisms have not yet been
elucidated. The study of [FeFe] hydrogenases has been
impeded by the lack of an easily manipulated expression/
activation system capable of producing these complex and
extremely oxygen-sensitive enzymes. Here we show the
first expression of functional [FeFe] hydrogenases in an
Escherichia coli-based cell-free transcription/translation
system. We have produced and matured both algal and
bacterial hydrogenases using E. coli cell extracts containing
the HydG, HydE, and HydF proteins from Shewanella
oneidensis. The current system produces
22 mg/mL of
active protein, constituting
44% of the total protein
produced. Active protein yield is greatly enhanced by pre-
incubation of the maturation enzyme-containing extract
with inorganic iron and sulfur for reconstitution of the
[Fe-S] clusters in HydG, HydE, and HydF. The absencegenases (Adams, 1990). Hydrogenases are especially im-
portant since they are key enzymes in many microbial
ecosystems (Spear et al., 2005) and may also be useful for
the biogenic production of hydrogen fuel as a renewable,
environmentally friendly resource. Hydrogenase enzymes
reversibly catalyze the condensation of two protons andher,1
1 North-South Mall, Stanford,
e-mail: jswartz@stanford.edu
pus Drive, Stanford,
0.1002/bit.21511
KEYWORDS: [FeFe] hydrogenases; cell-free protein synth-
esis; iron-sulfur proteins; maturation enzymes; hydrogen
production
Introduction
Many of the most important proteins in nature are difficult to
study because of complexities in their folding pathways
and final structures. Example enzymes with ecological
and industrial importance include nitrogenases (Rubio
and Ludden, 2005), methanogenic enzymes (Deppenmeier,

hydrogenase in vivo (Posewitz et al., 2004), though the
maturation mechanisms have not yet been elucidated.
Both [FeFe] hydrogenases and their maturation enzymes
This doubly heterologous system explored the functional
compatibility among components from multiple organisms.
This report represents the first expansion of cell extracts to
sequences or intra-operon structure. The resulting vectorare extremely sensitive to oxygen. Although the study and
engineering of complex proteins such as hydrogenases is
extremely important, progress has been hampered by the lack
of a convenient system capable of their expression and
activation.
Recombinant expression of [FeFe] hydrogenases in their
native hosts is possible but is difficult because these enzymes
occur in obligate anaerobes such as Clostridia or are
expressed in other bacteria and algae only under anaerobic
conditions. A recently developed Clostridium acetobutylicum
system yielded only 0.1 mg/L of purified recombinant algal
[FeFe] hydrogenase (Girbal et al., 2005). Recombinant
expression in Escherichia coli has been demonstrated, but
obtaining high yields of active protein is challenging and
requires coexpression of heterologous helper proteins
(Posewitz et al., 2004). The latest system, coexpressing
helper proteins from Clostridium acetobutylicum, is capable
of milligram per liter yields (King et al., 2006). For all in
vivo production scenarios, experimental control over the
expression and folding environment is extremely limited
due to the barrier imposed by the cell wall.
An alternative to in vivo protein production is cell-free
protein synthesis, in which protein expression is carried out
using cell extracts. Extracts from several organisms have
been used for cell-free protein synthesis (Endo and Sawasaki,
2003; Katzen et al., 2005; Tarui et al., 2000), but E. coli-based
systems are most common, and will be assumed hereafter.
Cell-free protein synthesis can generate high protein yields
(above 1 mg/mL) and preserves the procedural and genetic
advantages of working with E. coli.
Activation of complex proteins is one of the primary
challenges in a cell-free system, but, in contrast to in vivo
approaches, the absence of cell walls offers direct access to
the protein production and folding environment. This
allows facile addition of substrates, even to concentrations
unachievable in vivo. The ability to control the expression
environment has enabled production of proteins requiring
[Fe-S] clusters (Boyer et al., 2006), FAD cofactors (Knapp
and Swartz, 2004), heme groups (Miyazaki-Imamura et al.,
2003), and disulfide bonds (Yin and Swartz, 2004). Since
modern cell-free systems more closely mimic the natural
cytoplasmic folding environment of cells (Jewett and Swartz,
2004), protein folding can occur co-translationally, and
activation may be assisted by intact pathways from E. coli.
However, existing cell-free systems fail to activate complex
proteins requiring maturation enzymes unique to their
natural hosts.
We demonstrate the consistent capability of an E. coli-
based cell-free protein synthesis system to express and
mature [FeFe] hydrogenases at much higher levels than have
been achieved in vivo. Cell extract was prepared from cells
expressing maturation enzymes from Shewanella oneidensis
and was used to produce and activate hydrogenases from
Chlamydomonas reinhardtii and Clostridium pasteurianum.
60 Biotechnology and Bioengineering, Vol. 99, No. 1, January 1, 2008was transformed into E. coli strain BL21(DE3) and screened
for activity by coexpression with hydrogenase genes. The
helper protein plasmid from the colony that produced the
most active hydrogenase in coexpression experiments was
sequenced and found to be correct based on published data
(Accession #AE014299). This helper protein plasmid wasinclude heterologous maturation enzymes in a robust E. coli-
based platform and the first use of cell-free synthesis to study
oxygen-sensitive proteins. These advances will have parti-
cular impact on the production and study of other
metalloenzymes, anaerobic proteins, and proteins requiring
maturation enzymes.
Materials and Methods
Genes for CpI, HydA1, and HydGxEF
The gene and amino acid sequences for the CpI protein from
Clostridium pasteurianum were obtained from published
data (Accession #M81737). While preserving the amino acid
sequence, the gene was optimized for expression in E. coli by
adjusting codon usage to match that found in E. coli. In
addition, an NdeI restriction site was placed overlapping the
start codon and a SalI site was placed immediately after the
stop codon. A series of overlapping 60-mer oligonucleotides
was purchased commercially and used to synthesize the new
gene using published protocols (Stemmer et al., 1995). The
PCR product was digested with NdeI and SalI enzymes, and
ligated into the pK7 expression vector behind the standard
T7 promoter. The sequence was verified, and the plasmid
was named pK7sCpI.
The cDNA sequence encoding the HydA1 protein from
Chlamydomonas reinhardtii was obtained from published
data (Accession #AY055755) and truncated to remove bases
4–168, encoding the proposed signal sequence (Happe and
Kaminski, 2002). The gene was optimized, synthesized,
digested, and ligated into the pK7 vector as described above.
The resulting plasmid was labeled pK7sHydA1. For cell-free
expression experiments, purified plasmids were prepared
using Qiagen Maxiprep kits.
The maturation enzyme genes hydE, hydF, and hydG were
obtained from the gram-negative bacteria Shewanella
oneidensis. Genomic DNA was prepared using a Qiagen
genomic tip kit according to the manufacturer’s protocol.
The operon segment containing hydG, open reading frame
hydX, hydE, and hydF was amplified by PCR using primers
to insert an BspHI site overlapping the hydG start codon and
an AvrII site directly after the stop codon of hydF. This
segment was cloned in two steps into the pACYCduet vector
(Invitrogen, Carlsbad, CA) between NcoI and AvrII sites,
excising the second multiple cloning site. This placed the
genes behind a single T7 promoter without modifying geneDOI 10.1002/bit

named pACYC S.o. HydGxEF and was used for all anaerobic
cell-free production experiments.
with argon, stoppered, and shaken for 5 h. Cultures were
After completion of the fermentation, cells were harvested
using procedures designed to maintain the anaerobic nature
of the culture. Four-liter flasks were fitted with rubber
connected to the fermenter harvest port, and one of the
exhaust valves was opened. Cells were harvested through thetransferred into an anaerobic glove box (Coy Laboratories,
Grass Lake, MI) and pelleted by centrifugation, then
resuspended in permeabilization buffer (100 mM Tris/
HCl pH 8, 150 mM NaCl þ2% v/v Triton X-100). The
optical density of each suspension was measured and used to
normalize sample size to ensure equivalent cell content in
each assay. Background activity was measured using a
strain expressing S. oneidensis maturation enzymes and an
unrelated protein, b-lactamase.
For comparison, two plasmids used in previous coex-
pression experiments were kindly provided by M. Ghirardi,
one containing genes for the HydG and HydEF maturation
enzymes from C. reinhardtii (pACYC C.r. HydG, EF), and
one containing the unaltered hydA1 gene from C. reinhardtii
(pETHydA1; Posewitz et al., 2004).
The two plasmids containing genes for maturation
enzymes (pACYC C.r. HydG, EF and pACYC S.o. HydGxEF)
were transformed separately into E. coli strain BL21(DE3).
Resulting strains were further transformed with one of three
hydrogenase gene plasmids: pETHydA1, pK7sHydA1, or
pK7sCpI. Maturation enzymes and hydrogenase genes
were coexpressed in E. coli under anaerobic conditions
as described above. After coexpression, cells were pelletted
under anaerobic conditions and permeabilized by sus-
pension in anaerobic buffer containing 2% Triton X-100.
Hydrogenase activity was detected using a colorimetric assay
that couples hydrogenase activity with the reduction of
methyl viologen.
Culturing of Extract Source Cells
During the aerobic phase, cells were grown in a B. Braun
10 L fermentor in 8 L of defined media (see Supplemental
Table I), which included 10 g/L glucose and a mixture of
13 supplemented amino acids to promote rapid growth
(Zawada and Swartz, 2005). Successful extract preparations
came from fermentations with aerobic phase growth rates of

0.84 h1. Cells were still in the exponential growth phase
just before the anaerobic induction phase was started
(Fig. 3).In Vivo Coexpression of [FeFe] Hydrogenases and
Maturation Enzymes
S. oneidensis and C. reinhardtii maturation enzymes were
evaluated by coexpression with hydrogenases in E. coli. Fresh
overnight cultures of E. coli cells harboring maturation
enzyme and hydrogenase gene plasmids were subcultured in
5 mL of fresh LB media and grown aerobically at 30C to an
OD (600 nm) of 0.5–0.7. Protein production was induced by
addition of IPTG to 0.5 mM. Cells were grown aerobically
for one additional hour. Each culture tube was then flushedExtract Preparation
Cells were thawed in the anaerobic glove box and
resuspended in 1 mL S30 buffer per gram wet cells. Cell
clumps were eliminated by agitation and pipetting. Cells
were broken in a single pass through a high-pressure
homogenizer (>20,000 psi) inside the anaerobic glove box.
Cell lysate was immediately centrifuged twice for 30 min
at 30,000g at 4C also inside the anaerobic glove box.
Supernatant was removed, aliquotted, sealed in stoppered
anaerobic vials, frozen with liquid N2, and stored at 80C.
The incubation and dialysis steps that are commonly used in
extract preparation were eliminated because they were not
observed to increase, and in fact often decreased, the potency
of these anaerobic extracts for production of active [FeFe]
hydrogenases. Extracts were stable for several months.
Cell-Free Protein Production
PANOx-SP substrate mixtures (Jewett and Swartz, 2004)
were prepared with the following components listed with
their final concentrations: magnesium glutamate 8 mM;
ammonium glutamate 10 mM; potassium glutamate
175 mM; ATP 1.2 mM; GTP, UTP, and CTP 0.90 mM;
folinic acid 34 mg/mL; E. coli tRNA mixture 170 mg/mL;
20 amino acids 2 mM each; PEP 30 mM; NAD 0.33 mM;
Boyer et al.: Cell-Free Hydrogenase Synthesis 61cooling coil and into flasks while maintaining constant
purging with argon or nitrogen. After filling, all valves were
closed and the flasks were moved into an anaerobic glove
box where the contents were sealed in centrifuge bottles. All
further steps occurred either in the anaerobic glove box or in
centrifuge bottles filled and sealed in the glove box. After
centrifugation for 15 min at 6,000g, centrifuge bottles were
returned to the anaerobic glove box, the supernatant
was decanted, and cells were washed in S30 buffer (10 mM
Tris acetate pH 8.2, 14 mM magnesium acetate, 60 mM
potassium acetate, 2 mM DTT). Cells were again pelletted
and then transferred to stoppered anaerobic glass vials that
were sealed with aluminum crimp caps. Cell pellets were
frozen with liquid N2 and stored at 80C until extract was
prepared.stoppers, each with three inlet/outlet ports controlled by
valves: one for cell culture entry, one for entry of purge gas,
and one for exit of purge gas. The purge gas entry port was
connected to a length of silicon tubing that reached close to
the bottom of the flask so that argon or nitrogen could be
used to purge the flask from the bottom up. Flasks were
connected in series after a steel cooling coil immersed in an
ice water bath, and the system was purged with argon or
nitrogen before harvest began. The cooling coil was thenBiotechnology and Bioengineering. DOI 10.1002/bit

CoA 0.27 mM; oxalic acid 2.7 mM; putrescine 1 mM;
spermidine 1.5 mM; and ferrous ammonium sulfate
0.15 mM. After preparing the substrate mixture at 3.5
1 hydrogen analyzer (Peak Laboratories, Mountain View,
CA) under conditions for which the specific activity of
hydrogenase 1 from C. reinhardtii has been published
assay was quantified using a cell-free reaction mixture that
produced the unrelated protein chloramphenicol acetyl
produce active hydrogenases using conventional in vivofinal concentration in air, it was moved into an anaerobic
glove box, and the redox indicator resazurin was added to
1.75 mg/L. The solution was titrated with the reductant
Ti(III)citrate until a clear color persisted, then flash frozen
and stored at 80C in sealed glass vials.
For cell-free reactions, the following components were
added to the substrate mixture: plasmid DNA template 0.027
mg/mL, T7 RNA polymerase 0.1 mg/mL, S-adenosyl
methionine (SAM) 2 mM, 14C leucine 8.5 mM, cell extract
0.25 fraction of total reaction volume, and water. T7 RNA
polymerase was provided in a partially purified form after
overexpression in E. coli (Grodberg and Dunn, 1988). Total
protein produced in cell-free reactions was measured by
incorporation of radioactive leucine (Kim and Swartz, 2001).
Colorometric Hydrogen Consumption Activity Assay
Hydrogenase activity is commonly measured using the
colorometric reagent methyl viologen. Hydrogenase
enzymes reversibly catalyze the transfer of electrons between
hydrogen and methyl viologen according to the reaction:
2MVox þH2 !
hydrogenase
2MVred þ 2Hþ (1)
The reduced form of methyl viologen is dark blue, while the
oxidized form is clear.
The assay solution contained 2 mM methyl viologen in
50 mM Tris/HCl pH 8 with 1 mg/L of the redox indicator
resazurin. In the anaerobic glove box, the buffer was titrated
with Ti(III)citrate until the solution remained slightly blue.
Cell-free reaction mixtures were diluted 1,000- to 1,600-fold
in methyl viologen buffer. Color change was measured in a
96-well plate with an assay volume of 200 mL. The assay
mixture turned dark blue as hydrogenase catalyzed the
reduction of methyl viologen by hydrogen present in the
glove box at
2.5%. The change in absorbance was
monitored at 578 nm by a microplate spectrophotometer.
Both the buffer and the sample chamber of the instrument
were maintained at 37C. Background activity due to agents
in the cell extract was quantified using samples from a
parallel cell-free reaction mixture containing no DNA
template. Activity was estimated based on the initial slope of
absorbance versus time determined using linear regression.
In these highly diluted samples, the initial slope was
observed to be directly proportional to the amount of active
hydrogenase present. An extinction coefficient of 9.78 AU/
(mMcm) (Park et al., 1999) and a pathlength of 0.5 cm were
used to calculate the H2 consumption rate.
Gas Chromatograph-Based Hydrogen Production
Activity Assay
Hydrogen evolution from hydrogenase produced in the
cell-free system was quantified using a Peak Performer
62 Biotechnology and Bioengineering, Vol. 99, No. 1, January 1, 2008expression methods. E. coli does not contain genes for any
[FeFe] hydrogenases or their maturation enzymes and
cannot mature [FeFe] hydrogenases in an unmodified
strain (Voordouw et al., 1987). However, the first example of
expression of an active [FeFe] hydrogenase in E. coli was
recently accomplished by coexpression of the hydA1, hydEF,
and hydG genes from C. reinhardtii (Posewitz et al., 2004).
Subsequently, genetic stability and hydrogen production
were improved by coexpression of [FeFe] hydrogenases with
bacterial analogues of maturation enzymes (King et al.,
2006). In a parallel vein, we hypothesized that our E. coli-
based system would be better complemented by maturation
enzymes from the gram-negative bacterium S. oneidensis
than by enzymes from the eukaryotic alga C. reinhardtii.transferase. Incorporation of radioactivity was used to
quantify hydrogenase protein in the sample. Triplicate
measurements were performed for each of four dilutions
typical of the range used for experimentation. Both activity
assays gave a linear response over the measured range
(Supplemental Fig. 1).
Results and Discussion
In Vivo Coexpression of [FeFe] Hydrogenases and
Maturation Enzymes
With our primary objective being the expression and
screening of [FeFe] hydrogenase mutants, we first sought to(Happe and Naber, 1993). 9.5 mL glass vials containing
1 mL assay solution (5 mM methyl viologen, 25 mM sodium
dithionite, and 50 mM Tris/HCl pH 6.8) were stoppered and
sealed with an aluminum crimp seal in an anaerobic glove
box and then flushed with oxygen-free argon to remove
hydrogen. Two, four, six, and eight microliters of a 10
diluted cell-free reaction mixture was injected into the
reaction vials and mixed. Hydrogen production proceeded
at room temperature (
25C). Periodically, 100 mL samples
of the gas above the reaction were removed and injected into
the chromatograph to measure hydrogen content. Peak area
associated with hydrogen gas was converted to hydrogen
content using a calibration curve constructed using premix-
ed hydrogen gas standards.
Comparison of the Hydrogen Consumption and
Hydrogen Production Assays
Results the hydrogen consumption and hydrogen produc-
tion assays were compared for samples from the same cell-
free reaction mixtures. The background activity for eachDOI 10.1002/bit

To test this hypothesis, we coexpressed maturation
enzymes from S. oneidensis with [FeFe] hydrogenase
enzymes in E. coli as described in the Materials and
Methods section. Hydrogenase activity was observed for
both a bacterial and an algal hydrogenase in coexpression
experiments with S. oneidensis helper genes (Fig. 1, bars
3–5). Without coexpression of maturation enzymes, no
hydrogenase activity was observed (Fig. 1, bar 1). Also, no
significant activity was observed with coexpression of
C. reinhardtii hydrogenase and helper genes (Fig. 1, bar
2). These observations indicated that S. oneidensis helper
genes were active in hydrogenase maturation and that they
were better suited to complement E. coli for expression of
hydrogenases than helper genes from the alga C. reinhardtii.
These in vivo experiments were important to establish
the utility of S. oneidensis maturation enzymes, but the
magnitude and repeatability of hydrogenase activity from
these cultures were poor. Other highly variable data
have been reported from similar in vivo coexpression
experiments (Posewitz et al., 2004). To circumvent the
disadvantages encountered using in vivo expression and to
provide a more flexible expression platform, we sought to
produce active [FeFe] hydrogenases using cell-free protein
synthesis.
Optimization of Cell Extracts for Cell-Free Expression of
Yang et al., 2004; Yin and Swartz, 2004). Two unprecedented
adaptations in extract preparation were implemented for
expression of active hydrogenase proteins: the inclusion of
heterologous maturation enzymes, and the preparation of
extracts under strictly anaerobic conditions. Maturation
enzymes were introduced by expression of the hydG, hydE,
and hydF genes from the organism S. oneidensis during the
growth of the extract culture. The process of cell extract
preparation and its use in cell-free protein expression is
depicted in Figure 2.
The E. coli strain BL21(DE3) harboring the pACYC S.o.
HydGxEF plasmid was used to create a cell-free extract
capable of translating and maturing [FeFe]hydrogenases.
Cell growth was divided into two phases: an aerobic growth
phase, and an anaerobic induction and incubation phase.
Two of the maturation enzymes necessary for [FeFe]
hydrogenase maturation belong to the radical SAM family of
proteins (Posewitz et al., 2004), which are often oxygen
sensitive (Fontecave et al., 2001). For this reason, we
anticipated that the HydG, HydE, and HydF maturation
enzymes would need to be produced in E. coli under
anaerobic conditions.Figure 1. In vivo expression of [FeFe] hydrogenases. Helper proteins from either
C. reinhardtii or S. oneidensis were coexpressed in E. coli with hydrogenase genes
from either C. reinhardtii or C. pasteurianum. Cells were permeabilized with Triton X-
100 and tested for hydrogenase activity using the methyl viologen assay. Hydrogenase
activity was greatest in strains containing maturation enzymes from S. oneidensis.[FeFe] Hydrogenases
Cell extracts have been improved through strain engineering
and modified preparation procedures to enhance the cell-
free production of many types of proteins (Boyer et al., 2006;
Knapp and Swartz, 2004; Miyazaki-Imamura et al., 2003;Figure 2. Cell-free system for the production of oxygen-sensitive [FeFe] hydro-
genase proteins. Two unprecedented adaptations were made to produce active [FeFe]
hydrogenases: (1) heterologous helper proteins were expressed during growth of cells
to be used in extract preparation, and (2) extracts were prepared anaerobically. (3)
Cell extract is activated by pre-incubation with iron and sulfide to enhance hydro-
genase maturation capacity. (4) Hydrogenase is produced in an anaerobic cell-free
reaction by addition of substrates, DNA template, and RNA polymerase. The produc-
tion platform can be used at small scale to optimize system variables or to express and
screen protein mutants. The system can also be scaled up for purification of the target
protein.
Boyer et al.: Cell-Free Hydrogenase Synthesis 63
Biotechnology and Bioengineering. DOI 10.1002/bit

temperature was reduced to 15.5C. The culture was then
incubated for an additional 18 h before harvest. Little or no
growth was observed during the anaerobic induction
provide other helper proteins, chaperones, and maturation
enzymes.Anaerobic conditions were created by purging the culture
vessel with argon instead of air, and maturation enzyme
production was induced by the addition of IPTG. Several
variations of the timing of these two changes were evaluated,
including simultaneous induction and argon purge, induc-
tion after 30 min of anaerobic adaptation, and 30 min
of aerobic induction followed by argon purge. Our most
active extracts to date, with respect to active HydA1 protein
yields, were produced when maturation enzyme production
was induced with 500 mM IPTG 30 min after argon
purge began. To increase the effectiveness of the anaerobic
induction period, ferrous ammonium citrate was added to
0.33 mg/mL as a source of iron. Fumarate was added to
Figure 3. Growth of cells used for extract preparation. Cells were grown on
defined media at 37C to OD
6 before shifting to the argon purge. After 30 min
of anaerobic adaptation, maturation enzyme production was induced by addition of
IPTG to 0.5 mM and temperature was reduced to 15.5C for an additional 18 h. Little
growth occurred during the anaerobic induction phase.10 mM as an alternative electron acceptor to stimulate
metabolism in the absence of oxygen. Cysteine was added as
a sulfur source, but millimolar concentrations were
observed to cause the formation of a dark precipitate, so
its concentration was reduced to 0.05 mM.
Cell extracts were tested in in vitro hydrogenase
expression experiments as described in Materials and
Methods, and the results guided optimization of the culture
protocol. Early cell extracts produced significant quantities
of hydrogenase polypeptide, but little activity was observed,
indicating that the maturation capacity of the extracts was
limiting the accumulation of active hydrogenase. In an effort
to increase the concentration of soluble maturation enzymes
in the extract, we extended the period of anaerobic
induction and lowered the temperature. For some proteins,
lower production temperatures favor the formation of
soluble, folded product by slowing aggregation kinetics
(Rubach et al., 2005). After the aerobic growth phase and a
30-min anaerobic adaptation phase were conducted at 37C,
maturation enzyme production was induced and the
the rapid and high-throughput quantification of active
hydrogenase content of cell-free reaction mixtures.
64 Biotechnology and Bioengineering, Vol. 99, No. 1, January 1, 2008Optimization of Cell-Free Reaction Conditions
The cell-free system provides both ready access to the
reaction environment and the ability to perform many
small-scale reactions quickly and in parallel. This combina-
tion allows many process and reaction variables to be
quickly optimized. The magnesium concentration of the
substrate mixture had a strong effect on the yield of active
HydA1 protein. Optimal levels varied between 8 and 12 mM
for individual extract preparations. Compared to 378C,
incubation at 278C decreased the initial rate of total and
active hydrogenase accumulation, and total protein yieldsQuantification of Active Hydrogenase With the
Methyl Viologen Reduction Assay
Hydrogenase activity was most easily measured by adding a
small amount of cell-free reaction product to a buffered
solution of oxidized methyl viologen and monitoring
the formation of blue color using a microplate spectro-
photometer inside the anaerobic glove box (see Materials
and Methods). However, the amount of active hydrogenase
present could not be quantified without a value for the
specific activity of the C. reinhardtii HydA1 under these
conditions.
The specific activity of this enzyme purified from its
native source has been reported as 0.935 mmol H2/(minmg)
using a slightly different assay (Happe and Naber, 1993). We
measured the activity of hydrogenase produced in a cell-free
reaction using both the convenient hydrogen consumption
assay and the literature-based hydrogen production assay
(see Materials and Methods). The ratio of the average
activity measured in the two assays was multiplied by the
reported specific activity to yield a calibrated specific activity
for the hydrogen consumption assay of 0.90 0.19
mmol H2/(minmg active protein). This value allowed forphase (Fig. 3). Higher hydrogenase activity was produced
in cell-free protein reactions using these extracts.
In the past, enzymatic deficiencies in cell-free systems
have been addressed by addition of purified enzyme
solutions (Kim and Swartz, 2004; Miyazaki-Imamura
et al., 2003). The approach taken here was different in that
maturation enzymes were expressed in the cells used to make
the extract for cell-free protein synthesis reactions. In this
way, a single fermentation and extract preparation was used
to create machinery for both the cell-free translation and the
maturation of the target protein without requiring any
purification steps. This approach could be extended toDOI 10.1002/bit

were decreased. However, higher yields of active protein
were realized due to a longer duration of active protein
accumulation (Fig. 4a). Lower temperatures may favor
maturation by slowing polypeptide production rate.
Alternatively, inhibitory side reactions may be slowed at
lower temperatures, extending the duration of maturation.
Supplementation with 2 mM S-adenosyl methionine (SAM)
increased accumulation of active HydA1 by 35% (Fig. 4b).
No effects were observed from supplementation with 2 mM
S-adenosyl-homocysteine, carbamoyl phosphate, or addi-
tional glycine or cysteine above the 2 mM already added to
the reaction mixture (data not shown). These results fail
to support (but do not disprove) a recently advanced
theory that the CO and CN ligands that are bound to the
H-cluster could be derived from glycine (Peters et al.,
2006). Interestingly, SAM supplementation was required in
neously incorporate at least one [4Fe-4S] cluster when
hydrogenases, which instead contain an external peptideFigure 4. a: Time course of total and active protein accumulation for sHydA1 in
the cell-free system. Incubation at 37C increases initial accumulation rate for total and
active protein but results in lower overall yields of active protein. Error bars reflect the
standard deviation of five separate experiments. b: Effects of process variables on
total and active sHydA1 yields in the cell free system. Reconstitution of cell-free
extract and supplementation with SAM both have strong positive effects on accu-
mulation of active protein. The addition of iron and sulfide at reaction initiation is not
beneficial. Effects of reaction variables are less pronounced for total protein. Error
bars reflect the standard deviation of five separate experiments.loop that is hypothesized to be an alternative ferredoxin
interaction domain (Melis and Happe, 2001). Genes for
both the bacterial and algal type hydrogenases were
expressed in the cell-free system using the same reagents,
and in both cases, active enzyme was formed (Table I). This
Boyer et al.: Cell-Free Hydrogenase Synthesis 65incubated with inorganic iron and sulfide (Rubach
et al., 2005). HydF will incorporate a [4Fe-4S] cluster when
incubated with inorganic iron, cysteine, and cysteine
desulfurase (Brazzolotto et al., 2006). We hypothesized
that the maturation capacity of our extracts could be limited
if some of the maturation enzymes lacked their [Fe-S]
clusters. Cell extracts containing maturation enzymes were
incubated anaerobically with 1 mM each of DTT, ferrous
ammonium sulfate, and sodium sulfide for 2 h at glove
box temperature (
27C) before use in a cell-free protein
synthesis reaction. The formation of insoluble FeS did not
inhibit active hydrogenase accumulation. The anaerobic
pre-incubation resulted in a>65% increase in active protein
accumulation relative to reactions with untreated extract or
extract supplemented with iron and sulfide at reaction
initiation (Fig. 4b). These observations are consistent with
the involvement of the [Fe-S] clusters of the maturation
enzymes in hydrogenase activation and suggest that the
maturation enzymes produced in E. coli were originally only
partially activated. This anaerobic incubation is referred to
as the reconstitution procedure (Fig. 4).
Expression of Bacterial and Algal [FeFe] Hydrogenases
Bacterial and algal [FeFe] hydrogenases have highly
conserved domains surrounding the active site, but differ
markedly in other parts of the molecule. Structural studies of
the CpI bacterial hydrogenase reveal four [Fe-S] clusters
in N-terminal domains (Peters et al., 1998), which are
proposed to be involved in electron transfer from the natural
donor, ferredoxin. These clusters are absent in algalmillimolar concentrations, while active protein concentra-
tions were in the micromolar range. The reason this excess is
needed has not been determined but could be explained
if SAM is degraded through non-productive competing
pathways.
Reconstitution of Maturation Enzymes by
Pre-Incubation With Iron and Sulfide
The measure that most increased the yield of active
hydrogenase protein was an anaerobic pre-incubation of
the cell extract with inorganic iron and sulfide. HydG, HydE,
and HydF all contain [Fe-S] clusters that are likely involved
in maturation functions. Recent work has shown that the
maturation enzymes HydG and HydE will each sponta-Biotechnology and Bioengineering. DOI 10.1002/bit

the cell-free system’s lack of a cell wall offers superior control
other cofactors (Boyer et al., 2006; Knapp and Swartz, 2004),
membrane proteins, proteome libraries, and proteins
requiring chaperones (Yin and Swartz, 2004). In combina-
Katzen F, Chang G, Kudlicki W. 2005. The past, present and future of cell-
free protein synthesis. Trends Biotechnol 23(3):150–156.
Table I. Cell-free production of bacterial and Algal hydrogenases.
HydA1 CpIover the reaction environment, easier sampling, and the use
of various observation techniques. Since only one major
protein product is produced, the target is easily labeled by
incorporation of a radioactive or unnatural amino acid.
Reactions can be performed on small scale (15 mL) for
exploration of many variables or for screening of protein
mutants and then seamlessly scaled up (>500 mL) (Voloshin
and Swartz, 2005) for purification of target proteins.
Expression can be accomplished from plasmids or from
linear DNA templates (Michel-Reydellet et al., 2005) such as
might be created using mutagenesis methods.
The adaptations to the cell-free system described in
this work have enabled the production and maturation of
complex, oxygen-sensitive [FeFe] hydrogenases, which
represents a significant advance for the cell-free synthesis
platform. We expect that similar modifications to cell-free
systems will have direct applications for the study of otherimplies that the HydG, HydE, and HydF maturation
enzymes assist with maturation steps common to both
types of hydrogenases. About 40% of the produced HydA1
protein was activated in the cell-free system (Fig. 4b).
Interestingly, the simpler enzyme, HydA1, gave greater
activities in cell-free experiments, while its more complex
homolog CpI was more active in in vivo experiments. The
ability of the maturation enzymes to mature both types
of [FeFe] hydrogenases underscores their flexibility and
suggests that they promote the formation and installation of
the central H-cluster active site, since this is common to all
[FeFe] hydrogenases. Furthermore, the observed turnover
numbers for the cell-free produced enzymes of approxi-
mately 20,000 min1 (Table I) underscore the potential
importance of these Fe–Fe hydrogenases for hydrogen
production using genetically engineered organisms.
Conclusion
The cell-free system described in this work offers many
advantages for the production of [FeFe] hydrogenases and
other complex proteins. Compared to in vivo approaches,
Total protein (mg/mL cell-free reaction) 45 3 12 3
Hydrogenase activity
(nmol H2/(minmL cell-free reaction))
20 2 3 0.2
Hydrogenase specific activity
(mmol H2/(minnmol total protein))
21 2 16 4metalloproteins, oxygen-sensitive proteins, and proteins
requiring helpers to mature. Additionally, the cell-free
system has proven to be adaptable for the production of
mammalian proteins (Yang et al., 2005), proteins requiring
disulfide bonds (Yin and Swartz, 2004), proteins requiring
multiple subunits, proteins requiring [Fe-S] centers and
66 Biotechnology and Bioengineering, Vol. 99, No. 1, January 1, 2008Kim DM, Swartz JR. 2001. Regeneration of adenosine triphosphate from
glycolytic intermediates for cell-free protein synthesis. Biotechnol
Bioeng 74(4):309–316.
Kim DM, Swartz JR. 2004. Efficient production of a bioactive, multiple
disulfide-bonded protein using modified extracts of Escherichia coli.
Biotechnol Bioeng 85(2):122–129.
King PW, Posewitz MC, Ghirardi ML, Seibert M. 2006. Functional studies
of [FeFe] hydrogenase maturation in an Escherichia coli biosynthetic
system. J Bacteriol 188(6):2163–2172.tion, these capabilities establish this system as a highly
adaptable platform for the production and study of many
complex proteins.
The authors would like to thank Professor Alfred Spormann for many
valuable discussions and Megan Hoarfrost for her assistance with this
work. This work was funded by the Global Climate and Energy Project
at Stanford University.
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