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Production of xylitol by recombinant microalgae
Journal of Biotechnology 165 (2013) 178–183
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Journal of Biotechnology journal homepage: www.elsevier.com/locate/jbiotec
Production of xylitol by recombinant microalgae Azadeh Pourmir, Samaneh Noor-Mohammadi, Tyler W. Johannes ∗ Department of Chemical Engineering, The University of Tulsa, 800 S. Tucker Drive, Tulsa, OK 74104, United States
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Article history: Received 15 February 2013 Received in revised form 5 April 2013 Accepted 9 April 2013 Available online 15 April 2013 Keywords: Chlamydomonas reinhardtii Microalgae Chloroplast Xylitol
a b s t r a c t Microalgae have received significant attention recently as a potential low-cost host for the production of next-generation biofuels and natural products. Here we show that the chloroplast genome of the eukaryotic green microalga Chlamydomonas reinhardtii can be genetically engineered to produce xylitol through the introduction of a gene encoding a xylose reductase (XR) from the fungi Neurospora crassa. Increased levels of heterologous protein accumulation and xylitol production were achieved by synthesizing the XR gene in the chloroplast codon bias and by driving expression of the codon-optimized XR gene using a 16S/atpA promoter/5 -UTR fusion. These results demonstrate the feasibility of engineering microalgae to produce xylitol, and show the importance of codon optimizing the XR gene and using the 16S/atpA promoter/5 -UTR fusion to express XR in the chloroplast of C. reinhardtii. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Xylitol is a five-carbon sugar alcohol that is used as an artificial sweetener in the food and confectionary industries. Xylitol is roughly as sweet as sucrose and can be used as a sucrose substitute for diabetics (Emodi, 1978). In a series of field studies, a daily intake of xylitol was found to reduce tooth decay for high risk children and adolescents (Makinen et al., 1981). Consumer products such as toothpaste, chewing gum, mouthwash, and nasal spray are increasingly using xylitol in their formulations. A chemical method, involving the hydrogenation of xylose using a Raney nickel catalyst, is generally used for the industrial production of xylitol; however, this method presents several safety and environmental concerns because it requires high pressures and involves a toxic catalyst (Aminoff et al., 1978). More recent studies have focused on developing a safer, more environmentally friendly microbial route for the production of xylitol. These studies have primarily focused on investigating natural xylose-fermenting yeasts such as Candida tropicalis and Candida parapsilosis, or genetically engineering yeast strains such as Saccharomyces cerevisiae (Moon et al., 2002). For instance, high conversion of xylose to xylitol (>95%) was obtained by transforming S. cerevisiae with the gene encoding the xylose reductase (XR) from Pichia stipitis (Hallborn et al., 1991). XR catalyzes the first step in dxylose metabolism, reducing xylose to xylitol, and using NAD(P)H as a cofactor. In addition to various yeast strains, the recombinant expression of XR to facilitate xylitol production has been reported in the bacterium Escherichia coli (Cirino et al., 2006), but to date,
∗ Corresponding author. Tel.: +1 918 631 2947. E-mail address: [email protected] (T.W. Johannes). 0168-1656/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jbiotec.2013.04.002
no research efforts have focused on using microalgae for xylitol production. Compared to bacterial and yeast strains, microalgae may offer an attractive alternative for producing xylitol based on its simple growth requirements, generally regarded as safe (GRAS) designation, and long history as a source of food and food additives. Microalgae are a diverse group of prokaryotic and eukaryotic photosynthetic microorganisms that have useful applications in the food, nutritional, cosmetic, pharmaceutical, and biofuels industries (Coragliotti et al., 2011; Olaizola, 2003). Algae offer an attractive platform for the production of a variety of bioproducts because of their rapid growth rate, cost effective culturing, genetic manipulability, and ease of scale-up (Rasala et al., 2010). Among microalgae strains, Chlamydomonas reinhardtii is a popular model algae used in the study of photosynthesis and flagella function. This singled-cell eukaryotic green alga is genetically well characterized and has a significant molecular toolkit. All three genomes (nuclear, chloroplast, and mitochondrial) of C. reinhardtii have been fully sequenced and transformation methods that target each of these genomes have been developed (Specht et al., 2010). The chloroplast genome, in particular, is an attractive target for genetic engineering because the chloroplast has been shown to support high levels of heterologous protein accumulation (as high as 2-20% of total soluble protein (TSP)) and foreign DNA can be stably integrated at a specific location within the chloroplast genome (Rasala and Mayfield, 2011). In this study, we investigated the potential of using the microalga C. reinhardtii as a platform for the production of xylitol. The chloroplast genome of C. reinhardtii was transformed with a gene encoding a XR from the filamentous fungus Neurospora crassa. Strategies involving codon-optimizing of XR gene and using a hybrid-fusion promoter were also tested in an effort to increase heterologous XR protein levels in the chloroplast of C. reinhardtii.
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2. Materials and methods 2.1. Strains and media C. reinhardtii strain 137c (mt+) was obtained from Chlamydomonas Center (Duke University, Durham, NC). The plasmid pRS426m-xylose and the yeast strain S. cerevisiae YSG50 were obtained from Huimin Zhao at the University of Illinois at UrbanaChampaign. All algal cultures were grown to late logarithmic phase (typically 5-7 days) in tris-acetate-phosphate (TAP) medium containing 100 g/ml kanamycin under fluorescent white light (80 mol m−2 s−1 as measured by a Field Scout Quantum Meter (Plainfield, IL)) at 23 ◦ C on a rotary shaker at 100 rpm. E. coli DH5␣ was used for recombinant DNA manipulations and E. coli BL21 (DE3) was used as an expression host. S. cerevisiae YSG50 strain was grown in yeast extract–peptone–dextrose plus adenine (YPAD) medium at 30 ◦ C. 2.2. Plasmid construction Codon optimization of the XR gene was performed using software developed in-house. This program was written in the C++ programming language and optimizes a target gene sequence by substituting the most frequently used C. reinhardtii chloroplast codons (data obtained from http://www.kazusa.or.jp/codon (Nakamura et al., 2000)). The coding sequence of the codon optimized XR gene was ordered from Integrated DNA Technologies (Coralville, IA, USA) and designated optXR. The optXR gene was cloned into the NdeI/HindIII sites of the pET26b expression vector. The plasmids pTJ322-XR, pTJ322-optXR, and pTJ322-16S/optXR were constructed using a method as described by NoorMohammadi et al. (2012). The primers used to construct the three plasmids are shown in supplementary table 1. The 5 UTRs (psbA, 251 bp; atpA, 556 bp), 3 UTRs (psbA, 400 bp; atpA, 400 bp) and gene aphA6 (780 bp) were amplified via PCR from the plasmid pTJ322-aphA6-aadA. The XR gene (969 bp) was amplified from the plasmid pRS426m-xylose. The 16S rRNA sequence (Genbank accession number X03269.1; 219 bp) was amplified from chloroplast genomic DNA isolated from C. reinhardtii using a Wizard Genomic DNA isolation kit (Promega, Madison, WI, USA). The constructed plasmids were then subjected to restriction digestion analysis as described by Noor-Mohammadi et al. (2012). 2.3. Chloroplast transformations For chloroplast transformations, a 50 ml culture of C. reinhardtii was grown to late logarithmic phase in the presence of 0.5 mM 5 fluoro-2 -deoxyuridine (Sigma-Aldrich, St. Louis, MO, USA). Cells were harvested by centrifugation and resuspended in 4 ml of TAP medium. These cells were then spread onto TAP/agar plates containing 100 g/ml kanamycin. Chloroplast transformations were performed by particle bombardment (PDS-1000/He; Bio-Rad, Hercules, CA, USA) using DNA-coated gold particles (S550d; Seashell Technologies, San Diego, CA, USA). Particle bombardment parameters for the PDS-1000/He system were as follows: chamber vacuum of 28 in. Hg, helium pressure of 1350 psi, distance of 9 cm, and 3 mg of 0.55 m gold particles coated with 10 g of plasmid DNA. The particle-bombarded plates were placed under constant light for 1-2 weeks until transformed algae colonies appeared. Primary transformants were restreaked 4-5 times on TAP plates containing 100 l/ml kanamycin until homoplasmic lines were identified. 2.4. Protein expression, immunoblotting, and protein purification The plasmid pET26b-optXR was transformed into E. coli strain BL21 (DE3). Transformed cells were grown in LB media with
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shaking at 37 ◦ C. XR protein expression was induced by adding 0.5 mM isopropyl B-D-thiogalactopyranoside (IPTG) and placing the culture in a 30 ◦ C shaker until an OD600 of ∼0.6 was achieved. The cells were harvested by centrifugation, resuspended in 20 mM Tris–HCl pH 8.0, then placed in a −80 ◦ C freezer. After 1 h, the cells were thawed at room temperature, the cellular debris was removed by centrifugation, and the resulting supernatant was used to isolate the XR protein by affinity gel purification. Purification of FLAG-tagged XR protein was carried out using EZviewTM Red ANTI-FLAG® M2 Affinity Gel (Sigma, St. Louis, MO) according to the manufacturer’s instructions. Algal cell cultures were grown to late log phase in TAP media. Cells were harvested by centrifugation, and resuspended in 1 ml 20 mM Tris–HCl pH 8.0. The resuspended cells were lysed by sonication using a Misonix sonicator (Farmingdale, NY) with the amplitude set at 30% and with a pulse sequence of 7 s on and 10 s off, for 5 min. Samples were then centrifuged at 14,000 × g for 30 min at 4 ◦ C and the resulting supernatant used in western blot analysis. The protein concentration of the total soluble protein was measured using the BioRad Protein Reagent (Bio-Rad, Hercules, CA, USA) with bovine serum albumin as a standard. Approximately 3 g total soluble protein from each sample was separated on a 12% sodium dodecyl sulfate polyacrylamide gel (SDS–PAGE) and then transferred onto a methanol-treated PVDF (polyvinylidene fluoride, Millipore, Billerica, MA, USA) membrane. The PVDF membrane was blocked with 5% milk in TBST buffer (20 mM Tris, 150 mM NaCl, 0.02% Tween 20, pH 7.5) for 1 h at room temperature. The blocked membrane was incubated at 4 ◦ C overnight in TBST solution containing 1:1000 dilution of a rabbit anti-FLAG primary antibody (Cell Signaling Technology, Danvers, MA, USA), washed three times with TBST for 5 min, incubated with 1:2000 dilution of a horseradish peroxidase linked rabbit anti-rabbit IgG secondary antibody (Cell Signaling Technology, Danvers, MA, USA) for 1 h, then the membrane was washed with TBST for 30 min. A working solution of Pierce ECL Substrate (Thermo Scientific, Rockford, IL, USA) was prepared according to the manufacturer’s instructions and added to the membrane for 1 min. The membrane was removed from the substrate and analyzed by an Alpha Innotech FluorChem® HD2 imager and the software AlphaEaseFCTM . 2.5. Quantitative RT-PCR analysis of mRNA accumulation Cells were grown in TAP media under light until late log phase and then harvested by centrifugation at 4000 × g. Total RNA was isolated from the harvested cells using the Plant RNA Reagent Kit (Invitrogen, Carlsbad, CA, USA) according to manufacturer’s instructions. The concentration of extracted RNA was measured using a Thermo Scientific NanoDrop 2000c spectrophotometer and the integrity of the RNA was checked by agarose gel electrophoresis. Ten micrograms of total RNA was treated with DNase (Ambion Turbo DNA-free, Austin, TX, USA) to remove any contaminating genomic DNA. A Bio-Rad’s iScript cDNA Synthesis kit was used to synthesize cDNA from 1 g of DNase-treated total RNA. The synthesized cDNA was diluted 4-fold for use in the qPCRs. Each 20 l qPCR contained 10 l Bio-Rad iQTM SYBR Green Supermix, 0.5 M oligonucleotides, and 2 l diluted cDNA. Real-time qPCR was performed in triplicate for each sample using a StepOnePlus Real-Time PCR system (Applied Biosystems, Carlsbad, CA, USA). Thermocycling conditions consisted of a two-step sequence with an annealing/extension temperature of 60 ◦ C, followed by a melt curve to monitor for primer dimers. “No reverse transcriptase” for each sample was performed to monitor for possible genomic DNA contamination and “no template controls” for each primer pair were also performed in triplicate. The endogenous chloroplast gene rbcL of C. reinhardtii was used as a reference gene. Standard curves were generated for the heterologous genes XR, optXR, and the
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Fig. 1. Comparison of the XR and optXR coding regions. The amino acid sequence is shown below the nucleotide sequence. Codons that were changed are boxed.
reference gene rbcL (supplementary Fig. 6). Relative mRNA levels are expressed as a ratio of gene of interest to the rbcL gene. 2.6. Determination of xylitol production by HPLC To determine xylitol production, 50 ml cultures of wild-type C. reinhardtii and the three engineered C. reinhardtii strains were grown in TAP media under light until late log phase and then 5 ml of culture was harvested by centrifugation at 4000 × g for 5 min. The supernatant was removed and the cells were resuspended in 20 mM Tris–HCl pH 8.0 and 50 mM d-xylose and then incubated
at 25 ◦ C for 72 h. The incubated samples were mixed every 12 h by pipetting up and down several times. Samples were centrifuged and the supernatants were analyzed by high-performance liquid chromatography (HPLC). HPLC was performed using an Agilent 1260 Infinity Quaternary LC System (Agilent Technologies, Palo Alto, CA, USA) equipped with an online degasser and autosampler. d-Xylose and xylitol were separated using an Aminex HPX-87H (300 mm × 7.8 mm) ion exchange column (Bio-Rad, Hercules, CA, USA) and detected by a refractive index detector (RID). Substrate and product were separated using an isocratic elution of 5 mM sulfuric acid at a flow rate of 0.6 ml/min at 45 ◦ C. Retention times were
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Fig. 2. Three constructs used for XR expression in the chloroplasts of C. reinhardtii. Each construct contains a kanamycin-resistance gene aphA6 under the control of the psbA promoter and 5 -untranslated region (UTR) for selection purposes. All of the constructs include a 3× FLAG-tag sequence (DYKDDDDKS) fused to the C-terminus of the recombinant XR for Western blot analysis.
found to be 9.3 min for d-xylose and 10.7 min for xylitol. Each reaction was performed and analyzed in triplicate and the reported values are the average of these measurements with the associated standard deviation. 3. Results 3.1. Construction of the xylose reductase expression cassettes A well characterized xylose reductase (XR) from the fungus N. crassa was selected for this work based on its high activity, stability, and activity over a wide pH range (Woodyer et al., 2005). The C. reinhardtii chloroplast genome has a high AT content and therefore displays a strong codon bias (Morton, 1996). To improve protein expression, the xylose reductase gene sequence from N. crassa was optimized to match the codon usage of the C. reinhardtii chloroplast genome. The resulting codon-optimized xylose reductase gene (optXR) has a codon adaptive index (CAI) of 1.0 compared to a CAI of 0.2 for the native sequence (Fig. 1). Previous studies have demonstrated that different promoters and 5 -untranslated region (UTR) have a dramatic effect on mRNA and protein accumulation. In this study, we selected the atpA promoter and 5 -UTR and a 16S/atpA promoter/5 -UTR fusion, both of which have been shown to yield high levels of protein expression in the chloroplast of C. reinhardtii (Barnes et al., 2005; Rasala et al., 2011). We constructed three different expression cassettes to test whether a functional xylose reductase could be expressed in the chloroplast of C. reinhardtii. The three expression cassettes consisted of the XR gene under the control of the native atpA promoter and 5 -UTR, the optXR gene under the control of the atpA promoter and 5 -UTR, and the optXR gene under the control of the 16S/atpA promoter/5 -UTR fusion (Fig. 2). These three constructs were assembled into the plasmid pTJ322 using a method recently described by Noor-Mohammadi et al. (2012) (supplementary Fig. 1). For selection purposes, all three cassettes contained a kanamycin-resistance gene, aphA6, under the control of the psbA promoter and 5 -UTR. A 3× FLAG-tag epitope was added to the C-terminal of each protein sequence to allow detection by Western blot. Correct assembly of the three plasmids was confirmed by restriction digestion analysis (supplementary Fig. 2). 3.2. C. reinhardtii chloroplast transformation The constructed plasmids were transformed into the chloroplast genome of C. reinhardtii by particle gun bombardment. Primary transformants were selected on media supplemented with the antibiotic kanamycin and screened for integration and homoplasmicity by PCR. Each of the three constructs was stably integrated into the chloroplast genome downstream of the psbA gene and
Fig. 3. Analysis of the mRNA levels for the three transgenic strains of C. reinhardtii. mRNA levels are shown relative to the endogenous C. reinhardtii rbcL chloroplast gene. Data points represent averages of triplicate measurements with error bars representing ±1 standard deviation (SD).
upstream of the 5s rRNA region. Additional rounds of streaking for single colonies under selective pressure resulted in homoplasmic cell lines, in which all copies of the chloroplast genome contained the recombinant DNA. Homoplasmic strains were identified for all three constructs (supplementary Fig. 4). The transformation efficiency (colony forming unit per g of plasmid DNA) of the construct containing the 16S/atpA promoter/5 -UTR fusion was much lower than the transformation efficiency of the constructs containing the atpA promoter and 5 -UTR (data not shown). 3.3. Accumulation of mRNA in the transgenic strains Reverse transcriptase quantitative PCR (RT-qPCR) was used to determine the level of recombinant mRNAs for each of the three transgenic strains. The results of the RT-qPCR analysis are shown in Fig. 3. XR and optXR mRNA levels were calculated as a fold change relative to the expression of the endogenous rbcL gene for each strain. The strain expressing the codon-optimized optXR gene showed 4.3-fold less mRNA accumulation than the strain expressing the native XR gene. The 16S/atpA-optXR strain showed 1.2- and 5.4-fold higher mRNA accumulation than the atpA-XR and atpAoptXR strains, respectively. 3.4. Analysis of XR protein accumulation in transgenic C. reinhardtii chloroplasts The strains were analyzed by Western blot to determine the levels of XR protein accumulation in the C. reinhardtii transgenic cell lines. Three micrograms of total soluble protein (TSP) from wild-type C. reinhardtii, atpA-XR, atpA-optXR, and 16S/atpA-optXR strains were separated by SDS–PAGE, then the gel was subjected to either Coomassie staining or Western blot analysis. Although no discernible XR bands were apparent on the Coomassie-stained gel, all three strains accumulated XR protein as determined by Western blotting (Fig. 4). No detectable XR signal was observed in the wildtype C. reinhardtii cells. Quantification of XR accumulation against a dilution series of purified XR revealed that the enzyme accumulated to approximately 0.03%, 0.75%, and 2.2% of TSP in the atpA-XR, atpAoptXR, and 16S/atpA-optXR strains, respectively. Thus, the highest level of protein accumulation was observed in the 16S/atpA-optXR strain, demonstrating the importance of codon optimizing the xylose reductase gene and using a strong promoter/UTR to drive heterologous protein expression.
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Fig. 4. Accumulation and quantification of recombinant XR protein in the three transgenic lines. (A) Coomassie-stained gel in which 3 g of total soluble protein, isolated from the strains indicated, was subjected to 12% SDS–PAGE. XR expressed in E. coli BL21 (DE3) from the plasmid pET26-optXR was included as a positive control. (B) Western blot analysis of chloroplast-expressed XR proteins transferred to the PVDF membrane and probed with an anti-FLAG antibody. Total soluble protein (3 g) from each of the three transgenic strains was compared against a dilution series of purified XR to quantify XR accumulation. Densitometric analysis of the immunoblot signals using the software AlphaEaseFCTM indicates that XR accumulates to approximately 2.2% of total soluble protein for the 16S/optXR transformant. WT C. reinhardtii lysate is included as a negative control.
3.5. Xylitol production in the transgenic C. reinhardtii strains
4. Discussion
Whole-cell feeding experiments with xylose were performed to determine xylitol production in the C. reinhardtii transgenic cell lines. Algal cell cultures were grown to late log phase (5-7 days) in TAP media, harvested by centrifugation, then resuspended in a buffered solution containing 50 mM d-xylose. Resting cells were incubated at 25 ◦ C for 72 h and analyzed by HPLC. The results of the HPLC analysis are shown in Fig. 5. No xylitol was detected in the wild-type C. reinhardtii cells. The C. reinhardtii strains atpA-XR, atpA-optXR, and 16S/atpA-optXR were found to produce 0.04 g/L, 0.27 g/L, and 0.38 g/L of xylitol, respectively. For all three transgenic strains, all the unreacted xylose was determined to be in the wholecell reaction media. Although the 16S/atpA-optXR strain produced the highest amount of xylitol, its overall conversion was relatively low at 5.1%.
In this study, we have shown that a functional xylose reductase (XR) from N. crassa can be heterologously expressed in the chloroplast of C. reinhardtii. We assembled and integrated three different expression cassettes containing XR into the chloroplast genome of C. reinhardtii. Protein expression, mRNA levels, and xylitol production were quantitatively compared for each of the transgenic strains expressing these cassettes. Several factors have been identified that affect heterologous protein expression in the chloroplast of C. reinhardtii. The C. reinhardtii chloroplast genome displays a strong codon bias, with A or T (80%) preferred at the third position, and thus codon optimization of the target transgene has been shown to significantly improve protein yields (Franklin et al., 2002; Yoon et al., 2011). This conclusion is supported by our results showing that the C. reinhardtii chloroplasts transformed with optXR accumulated 25-fold more XR protein than native XR gene under the same atpA promoter/5 -UTR. Despite accumulating more XR protein, the strain expressing the optXR gene accumulated 4.3-fold less mRNA than the strain expressing XR. Previous studies have shown that the accumulation of chimeric mRNA in the chloroplast of C. reinhardtii is generally proportional to protein accumulation; however, some notable exceptions have been observed (Barnes et al., 2005), and thus this result appears to be one of these exceptions. Another factor that has been shown to dramatically influence the expression of heterologous proteins in the chloroplast of C. reinhardtii is the selection of the promoter and 5 -UTR used to drive transgene expression. The atpA promoter/5 -UTR was selected for this work based on its ability to produce high levels of chimeric mRNA and recombinant protein in previous studies (Barnes et al., 2005; Noor-Mohammadi et al., 2012). Recently, the fusion of the 16S promoter to the atpA 5 -UTR was shown to boost mRNA and heterologous protein to even higher levels in the chloroplast of C. reinhardtii (Rasala et al., 2011). A similar effect was observed in this work by fusing the 16S promoter to the atpA 5 -UTR to drive optXR expression. The 16S/atpA-optXR strain displayed 5.4-fold higher mRNA levels and 2.9-fold higher protein accumulation than the atpA-optXR strain.
Fig. 5. Comparison of xylitol production from wild-type (WT) C. reinhardtii and the three transgenic strains. No xylitol production was observed in the WT strain. SD of the triplicate measurements is shown in the error bars.
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Although we were able to achieve xylitol production in C. reinhardtii, the 0.05 g/g (xylitol/xylose) yield we obtained using the 16S/atpA-optXR strain was relatively low compared to other xylitol producing microorganisms. For instance, Lee and coworkers achieved a xylitol production yield of 0.92 g/g (xylitol/xylose) by feeding xylose to an engineered strain of C. tropicalis (Lee et al., 2003). Two possible explanations may account for this low yield. First, it is possible that xylose may not be transported efficiently into the chloroplast of C. reinhardtii. The transport of xylose has been shown to be a rate-limiting step for xylose metabolism in other organisms such as the yeast strain S. cerevisiae (Gardonyi et al., 2003). For xylose to be transported into the stroma of the chloroplast of C. reinhardtii, it must pass through multiple membranes including the plasma membrane and the inner and outer membranes of the chloroplast. Passive diffusion through each of these membranes more than likely accounts for the small amount of conversion observed in this study. The outer membrane of C. reinhardtii also does not appear to contain any xylose transporters based on an analysis of its genome sequence. It is possible that the additional expression of xylose transporters in C. reinhardtii would result in higher xylitol yields. Sugar transporters have been successfully introduced into several microalgae strains, including C. reinhardtii, to facilitate the transfer of glucose across the outer membrane and enable heterotrophic growth (Doebbe et al., 2007). Another possible explanation is that there may not be a sufficient amount of the NAD(P)H cofactor available in the chloroplast of C. reinhardtii. The XR from N. crassa uses both NADPH and NADH, thus the depletion of these factors could result in a cofactor imbalance in the chloroplast. Cofactor imbalances have been shown to be a major factor in controlling xylitol accumulation in several yeast strains (van Dijken and Scheffers, 1986; Young et al., 2010). 5. Conclusions In this work, we show that xylitol production in C. reinhardtii is feasible by genetically modifying the chloroplast genome. Despite achieving xylitol production, the overall yield (0.05 g xylitol/g xylose) from C. reinhardtii was relatively low. Further yield improvements are therefore necessary to make the engineered C. reinhardtii strain competitive with other xylitol producing microorganisms. Another possible route for enabling xylitol production in C. reinhardtii would be to target the nuclear genome. Targeting the nuclear genome may alleviate some of the issues involved with transporting xylose into the chloroplast; however, this approach is likely to run into difficulties because transgene silencing remains a significant obstacle to the expression of recombinant proteins in the nucleus of C. reinhardtii (Specht et al., 2010). Acknowledgments This work was supported by NSF EPSCoR award EPS0814361 to the State of Oklahoma and NSF award CHE1048784. A.P. and S.N. were supported by Conoco-Phillips fellowships. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jbiotec. 2013.04.002.
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Journal of Biotechnology journal homepage: www.elsevier.com/locate/jbiotec
Production of xylitol by recombinant microalgae Azadeh Pourmir, Samaneh Noor-Mohammadi, Tyler W. Johannes ∗ Department of Chemical Engineering, The University of Tulsa, 800 S. Tucker Drive, Tulsa, OK 74104, United States
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Article history: Received 15 February 2013 Received in revised form 5 April 2013 Accepted 9 April 2013 Available online 15 April 2013 Keywords: Chlamydomonas reinhardtii Microalgae Chloroplast Xylitol
a b s t r a c t Microalgae have received significant attention recently as a potential low-cost host for the production of next-generation biofuels and natural products. Here we show that the chloroplast genome of the eukaryotic green microalga Chlamydomonas reinhardtii can be genetically engineered to produce xylitol through the introduction of a gene encoding a xylose reductase (XR) from the fungi Neurospora crassa. Increased levels of heterologous protein accumulation and xylitol production were achieved by synthesizing the XR gene in the chloroplast codon bias and by driving expression of the codon-optimized XR gene using a 16S/atpA promoter/5 -UTR fusion. These results demonstrate the feasibility of engineering microalgae to produce xylitol, and show the importance of codon optimizing the XR gene and using the 16S/atpA promoter/5 -UTR fusion to express XR in the chloroplast of C. reinhardtii. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Xylitol is a five-carbon sugar alcohol that is used as an artificial sweetener in the food and confectionary industries. Xylitol is roughly as sweet as sucrose and can be used as a sucrose substitute for diabetics (Emodi, 1978). In a series of field studies, a daily intake of xylitol was found to reduce tooth decay for high risk children and adolescents (Makinen et al., 1981). Consumer products such as toothpaste, chewing gum, mouthwash, and nasal spray are increasingly using xylitol in their formulations. A chemical method, involving the hydrogenation of xylose using a Raney nickel catalyst, is generally used for the industrial production of xylitol; however, this method presents several safety and environmental concerns because it requires high pressures and involves a toxic catalyst (Aminoff et al., 1978). More recent studies have focused on developing a safer, more environmentally friendly microbial route for the production of xylitol. These studies have primarily focused on investigating natural xylose-fermenting yeasts such as Candida tropicalis and Candida parapsilosis, or genetically engineering yeast strains such as Saccharomyces cerevisiae (Moon et al., 2002). For instance, high conversion of xylose to xylitol (>95%) was obtained by transforming S. cerevisiae with the gene encoding the xylose reductase (XR) from Pichia stipitis (Hallborn et al., 1991). XR catalyzes the first step in dxylose metabolism, reducing xylose to xylitol, and using NAD(P)H as a cofactor. In addition to various yeast strains, the recombinant expression of XR to facilitate xylitol production has been reported in the bacterium Escherichia coli (Cirino et al., 2006), but to date,
∗ Corresponding author. Tel.: +1 918 631 2947. E-mail address: [email protected] (T.W. Johannes). 0168-1656/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jbiotec.2013.04.002
no research efforts have focused on using microalgae for xylitol production. Compared to bacterial and yeast strains, microalgae may offer an attractive alternative for producing xylitol based on its simple growth requirements, generally regarded as safe (GRAS) designation, and long history as a source of food and food additives. Microalgae are a diverse group of prokaryotic and eukaryotic photosynthetic microorganisms that have useful applications in the food, nutritional, cosmetic, pharmaceutical, and biofuels industries (Coragliotti et al., 2011; Olaizola, 2003). Algae offer an attractive platform for the production of a variety of bioproducts because of their rapid growth rate, cost effective culturing, genetic manipulability, and ease of scale-up (Rasala et al., 2010). Among microalgae strains, Chlamydomonas reinhardtii is a popular model algae used in the study of photosynthesis and flagella function. This singled-cell eukaryotic green alga is genetically well characterized and has a significant molecular toolkit. All three genomes (nuclear, chloroplast, and mitochondrial) of C. reinhardtii have been fully sequenced and transformation methods that target each of these genomes have been developed (Specht et al., 2010). The chloroplast genome, in particular, is an attractive target for genetic engineering because the chloroplast has been shown to support high levels of heterologous protein accumulation (as high as 2-20% of total soluble protein (TSP)) and foreign DNA can be stably integrated at a specific location within the chloroplast genome (Rasala and Mayfield, 2011). In this study, we investigated the potential of using the microalga C. reinhardtii as a platform for the production of xylitol. The chloroplast genome of C. reinhardtii was transformed with a gene encoding a XR from the filamentous fungus Neurospora crassa. Strategies involving codon-optimizing of XR gene and using a hybrid-fusion promoter were also tested in an effort to increase heterologous XR protein levels in the chloroplast of C. reinhardtii.
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2. Materials and methods 2.1. Strains and media C. reinhardtii strain 137c (mt+) was obtained from Chlamydomonas Center (Duke University, Durham, NC). The plasmid pRS426m-xylose and the yeast strain S. cerevisiae YSG50 were obtained from Huimin Zhao at the University of Illinois at UrbanaChampaign. All algal cultures were grown to late logarithmic phase (typically 5-7 days) in tris-acetate-phosphate (TAP) medium containing 100 g/ml kanamycin under fluorescent white light (80 mol m−2 s−1 as measured by a Field Scout Quantum Meter (Plainfield, IL)) at 23 ◦ C on a rotary shaker at 100 rpm. E. coli DH5␣ was used for recombinant DNA manipulations and E. coli BL21 (DE3) was used as an expression host. S. cerevisiae YSG50 strain was grown in yeast extract–peptone–dextrose plus adenine (YPAD) medium at 30 ◦ C. 2.2. Plasmid construction Codon optimization of the XR gene was performed using software developed in-house. This program was written in the C++ programming language and optimizes a target gene sequence by substituting the most frequently used C. reinhardtii chloroplast codons (data obtained from http://www.kazusa.or.jp/codon (Nakamura et al., 2000)). The coding sequence of the codon optimized XR gene was ordered from Integrated DNA Technologies (Coralville, IA, USA) and designated optXR. The optXR gene was cloned into the NdeI/HindIII sites of the pET26b expression vector. The plasmids pTJ322-XR, pTJ322-optXR, and pTJ322-16S/optXR were constructed using a method as described by NoorMohammadi et al. (2012). The primers used to construct the three plasmids are shown in supplementary table 1. The 5 UTRs (psbA, 251 bp; atpA, 556 bp), 3 UTRs (psbA, 400 bp; atpA, 400 bp) and gene aphA6 (780 bp) were amplified via PCR from the plasmid pTJ322-aphA6-aadA. The XR gene (969 bp) was amplified from the plasmid pRS426m-xylose. The 16S rRNA sequence (Genbank accession number X03269.1; 219 bp) was amplified from chloroplast genomic DNA isolated from C. reinhardtii using a Wizard Genomic DNA isolation kit (Promega, Madison, WI, USA). The constructed plasmids were then subjected to restriction digestion analysis as described by Noor-Mohammadi et al. (2012). 2.3. Chloroplast transformations For chloroplast transformations, a 50 ml culture of C. reinhardtii was grown to late logarithmic phase in the presence of 0.5 mM 5 fluoro-2 -deoxyuridine (Sigma-Aldrich, St. Louis, MO, USA). Cells were harvested by centrifugation and resuspended in 4 ml of TAP medium. These cells were then spread onto TAP/agar plates containing 100 g/ml kanamycin. Chloroplast transformations were performed by particle bombardment (PDS-1000/He; Bio-Rad, Hercules, CA, USA) using DNA-coated gold particles (S550d; Seashell Technologies, San Diego, CA, USA). Particle bombardment parameters for the PDS-1000/He system were as follows: chamber vacuum of 28 in. Hg, helium pressure of 1350 psi, distance of 9 cm, and 3 mg of 0.55 m gold particles coated with 10 g of plasmid DNA. The particle-bombarded plates were placed under constant light for 1-2 weeks until transformed algae colonies appeared. Primary transformants were restreaked 4-5 times on TAP plates containing 100 l/ml kanamycin until homoplasmic lines were identified. 2.4. Protein expression, immunoblotting, and protein purification The plasmid pET26b-optXR was transformed into E. coli strain BL21 (DE3). Transformed cells were grown in LB media with
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shaking at 37 ◦ C. XR protein expression was induced by adding 0.5 mM isopropyl B-D-thiogalactopyranoside (IPTG) and placing the culture in a 30 ◦ C shaker until an OD600 of ∼0.6 was achieved. The cells were harvested by centrifugation, resuspended in 20 mM Tris–HCl pH 8.0, then placed in a −80 ◦ C freezer. After 1 h, the cells were thawed at room temperature, the cellular debris was removed by centrifugation, and the resulting supernatant was used to isolate the XR protein by affinity gel purification. Purification of FLAG-tagged XR protein was carried out using EZviewTM Red ANTI-FLAG® M2 Affinity Gel (Sigma, St. Louis, MO) according to the manufacturer’s instructions. Algal cell cultures were grown to late log phase in TAP media. Cells were harvested by centrifugation, and resuspended in 1 ml 20 mM Tris–HCl pH 8.0. The resuspended cells were lysed by sonication using a Misonix sonicator (Farmingdale, NY) with the amplitude set at 30% and with a pulse sequence of 7 s on and 10 s off, for 5 min. Samples were then centrifuged at 14,000 × g for 30 min at 4 ◦ C and the resulting supernatant used in western blot analysis. The protein concentration of the total soluble protein was measured using the BioRad Protein Reagent (Bio-Rad, Hercules, CA, USA) with bovine serum albumin as a standard. Approximately 3 g total soluble protein from each sample was separated on a 12% sodium dodecyl sulfate polyacrylamide gel (SDS–PAGE) and then transferred onto a methanol-treated PVDF (polyvinylidene fluoride, Millipore, Billerica, MA, USA) membrane. The PVDF membrane was blocked with 5% milk in TBST buffer (20 mM Tris, 150 mM NaCl, 0.02% Tween 20, pH 7.5) for 1 h at room temperature. The blocked membrane was incubated at 4 ◦ C overnight in TBST solution containing 1:1000 dilution of a rabbit anti-FLAG primary antibody (Cell Signaling Technology, Danvers, MA, USA), washed three times with TBST for 5 min, incubated with 1:2000 dilution of a horseradish peroxidase linked rabbit anti-rabbit IgG secondary antibody (Cell Signaling Technology, Danvers, MA, USA) for 1 h, then the membrane was washed with TBST for 30 min. A working solution of Pierce ECL Substrate (Thermo Scientific, Rockford, IL, USA) was prepared according to the manufacturer’s instructions and added to the membrane for 1 min. The membrane was removed from the substrate and analyzed by an Alpha Innotech FluorChem® HD2 imager and the software AlphaEaseFCTM . 2.5. Quantitative RT-PCR analysis of mRNA accumulation Cells were grown in TAP media under light until late log phase and then harvested by centrifugation at 4000 × g. Total RNA was isolated from the harvested cells using the Plant RNA Reagent Kit (Invitrogen, Carlsbad, CA, USA) according to manufacturer’s instructions. The concentration of extracted RNA was measured using a Thermo Scientific NanoDrop 2000c spectrophotometer and the integrity of the RNA was checked by agarose gel electrophoresis. Ten micrograms of total RNA was treated with DNase (Ambion Turbo DNA-free, Austin, TX, USA) to remove any contaminating genomic DNA. A Bio-Rad’s iScript cDNA Synthesis kit was used to synthesize cDNA from 1 g of DNase-treated total RNA. The synthesized cDNA was diluted 4-fold for use in the qPCRs. Each 20 l qPCR contained 10 l Bio-Rad iQTM SYBR Green Supermix, 0.5 M oligonucleotides, and 2 l diluted cDNA. Real-time qPCR was performed in triplicate for each sample using a StepOnePlus Real-Time PCR system (Applied Biosystems, Carlsbad, CA, USA). Thermocycling conditions consisted of a two-step sequence with an annealing/extension temperature of 60 ◦ C, followed by a melt curve to monitor for primer dimers. “No reverse transcriptase” for each sample was performed to monitor for possible genomic DNA contamination and “no template controls” for each primer pair were also performed in triplicate. The endogenous chloroplast gene rbcL of C. reinhardtii was used as a reference gene. Standard curves were generated for the heterologous genes XR, optXR, and the
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Fig. 1. Comparison of the XR and optXR coding regions. The amino acid sequence is shown below the nucleotide sequence. Codons that were changed are boxed.
reference gene rbcL (supplementary Fig. 6). Relative mRNA levels are expressed as a ratio of gene of interest to the rbcL gene. 2.6. Determination of xylitol production by HPLC To determine xylitol production, 50 ml cultures of wild-type C. reinhardtii and the three engineered C. reinhardtii strains were grown in TAP media under light until late log phase and then 5 ml of culture was harvested by centrifugation at 4000 × g for 5 min. The supernatant was removed and the cells were resuspended in 20 mM Tris–HCl pH 8.0 and 50 mM d-xylose and then incubated
at 25 ◦ C for 72 h. The incubated samples were mixed every 12 h by pipetting up and down several times. Samples were centrifuged and the supernatants were analyzed by high-performance liquid chromatography (HPLC). HPLC was performed using an Agilent 1260 Infinity Quaternary LC System (Agilent Technologies, Palo Alto, CA, USA) equipped with an online degasser and autosampler. d-Xylose and xylitol were separated using an Aminex HPX-87H (300 mm × 7.8 mm) ion exchange column (Bio-Rad, Hercules, CA, USA) and detected by a refractive index detector (RID). Substrate and product were separated using an isocratic elution of 5 mM sulfuric acid at a flow rate of 0.6 ml/min at 45 ◦ C. Retention times were
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Fig. 2. Three constructs used for XR expression in the chloroplasts of C. reinhardtii. Each construct contains a kanamycin-resistance gene aphA6 under the control of the psbA promoter and 5 -untranslated region (UTR) for selection purposes. All of the constructs include a 3× FLAG-tag sequence (DYKDDDDKS) fused to the C-terminus of the recombinant XR for Western blot analysis.
found to be 9.3 min for d-xylose and 10.7 min for xylitol. Each reaction was performed and analyzed in triplicate and the reported values are the average of these measurements with the associated standard deviation. 3. Results 3.1. Construction of the xylose reductase expression cassettes A well characterized xylose reductase (XR) from the fungus N. crassa was selected for this work based on its high activity, stability, and activity over a wide pH range (Woodyer et al., 2005). The C. reinhardtii chloroplast genome has a high AT content and therefore displays a strong codon bias (Morton, 1996). To improve protein expression, the xylose reductase gene sequence from N. crassa was optimized to match the codon usage of the C. reinhardtii chloroplast genome. The resulting codon-optimized xylose reductase gene (optXR) has a codon adaptive index (CAI) of 1.0 compared to a CAI of 0.2 for the native sequence (Fig. 1). Previous studies have demonstrated that different promoters and 5 -untranslated region (UTR) have a dramatic effect on mRNA and protein accumulation. In this study, we selected the atpA promoter and 5 -UTR and a 16S/atpA promoter/5 -UTR fusion, both of which have been shown to yield high levels of protein expression in the chloroplast of C. reinhardtii (Barnes et al., 2005; Rasala et al., 2011). We constructed three different expression cassettes to test whether a functional xylose reductase could be expressed in the chloroplast of C. reinhardtii. The three expression cassettes consisted of the XR gene under the control of the native atpA promoter and 5 -UTR, the optXR gene under the control of the atpA promoter and 5 -UTR, and the optXR gene under the control of the 16S/atpA promoter/5 -UTR fusion (Fig. 2). These three constructs were assembled into the plasmid pTJ322 using a method recently described by Noor-Mohammadi et al. (2012) (supplementary Fig. 1). For selection purposes, all three cassettes contained a kanamycin-resistance gene, aphA6, under the control of the psbA promoter and 5 -UTR. A 3× FLAG-tag epitope was added to the C-terminal of each protein sequence to allow detection by Western blot. Correct assembly of the three plasmids was confirmed by restriction digestion analysis (supplementary Fig. 2). 3.2. C. reinhardtii chloroplast transformation The constructed plasmids were transformed into the chloroplast genome of C. reinhardtii by particle gun bombardment. Primary transformants were selected on media supplemented with the antibiotic kanamycin and screened for integration and homoplasmicity by PCR. Each of the three constructs was stably integrated into the chloroplast genome downstream of the psbA gene and
Fig. 3. Analysis of the mRNA levels for the three transgenic strains of C. reinhardtii. mRNA levels are shown relative to the endogenous C. reinhardtii rbcL chloroplast gene. Data points represent averages of triplicate measurements with error bars representing ±1 standard deviation (SD).
upstream of the 5s rRNA region. Additional rounds of streaking for single colonies under selective pressure resulted in homoplasmic cell lines, in which all copies of the chloroplast genome contained the recombinant DNA. Homoplasmic strains were identified for all three constructs (supplementary Fig. 4). The transformation efficiency (colony forming unit per g of plasmid DNA) of the construct containing the 16S/atpA promoter/5 -UTR fusion was much lower than the transformation efficiency of the constructs containing the atpA promoter and 5 -UTR (data not shown). 3.3. Accumulation of mRNA in the transgenic strains Reverse transcriptase quantitative PCR (RT-qPCR) was used to determine the level of recombinant mRNAs for each of the three transgenic strains. The results of the RT-qPCR analysis are shown in Fig. 3. XR and optXR mRNA levels were calculated as a fold change relative to the expression of the endogenous rbcL gene for each strain. The strain expressing the codon-optimized optXR gene showed 4.3-fold less mRNA accumulation than the strain expressing the native XR gene. The 16S/atpA-optXR strain showed 1.2- and 5.4-fold higher mRNA accumulation than the atpA-XR and atpAoptXR strains, respectively. 3.4. Analysis of XR protein accumulation in transgenic C. reinhardtii chloroplasts The strains were analyzed by Western blot to determine the levels of XR protein accumulation in the C. reinhardtii transgenic cell lines. Three micrograms of total soluble protein (TSP) from wild-type C. reinhardtii, atpA-XR, atpA-optXR, and 16S/atpA-optXR strains were separated by SDS–PAGE, then the gel was subjected to either Coomassie staining or Western blot analysis. Although no discernible XR bands were apparent on the Coomassie-stained gel, all three strains accumulated XR protein as determined by Western blotting (Fig. 4). No detectable XR signal was observed in the wildtype C. reinhardtii cells. Quantification of XR accumulation against a dilution series of purified XR revealed that the enzyme accumulated to approximately 0.03%, 0.75%, and 2.2% of TSP in the atpA-XR, atpAoptXR, and 16S/atpA-optXR strains, respectively. Thus, the highest level of protein accumulation was observed in the 16S/atpA-optXR strain, demonstrating the importance of codon optimizing the xylose reductase gene and using a strong promoter/UTR to drive heterologous protein expression.
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Fig. 4. Accumulation and quantification of recombinant XR protein in the three transgenic lines. (A) Coomassie-stained gel in which 3 g of total soluble protein, isolated from the strains indicated, was subjected to 12% SDS–PAGE. XR expressed in E. coli BL21 (DE3) from the plasmid pET26-optXR was included as a positive control. (B) Western blot analysis of chloroplast-expressed XR proteins transferred to the PVDF membrane and probed with an anti-FLAG antibody. Total soluble protein (3 g) from each of the three transgenic strains was compared against a dilution series of purified XR to quantify XR accumulation. Densitometric analysis of the immunoblot signals using the software AlphaEaseFCTM indicates that XR accumulates to approximately 2.2% of total soluble protein for the 16S/optXR transformant. WT C. reinhardtii lysate is included as a negative control.
3.5. Xylitol production in the transgenic C. reinhardtii strains
4. Discussion
Whole-cell feeding experiments with xylose were performed to determine xylitol production in the C. reinhardtii transgenic cell lines. Algal cell cultures were grown to late log phase (5-7 days) in TAP media, harvested by centrifugation, then resuspended in a buffered solution containing 50 mM d-xylose. Resting cells were incubated at 25 ◦ C for 72 h and analyzed by HPLC. The results of the HPLC analysis are shown in Fig. 5. No xylitol was detected in the wild-type C. reinhardtii cells. The C. reinhardtii strains atpA-XR, atpA-optXR, and 16S/atpA-optXR were found to produce 0.04 g/L, 0.27 g/L, and 0.38 g/L of xylitol, respectively. For all three transgenic strains, all the unreacted xylose was determined to be in the wholecell reaction media. Although the 16S/atpA-optXR strain produced the highest amount of xylitol, its overall conversion was relatively low at 5.1%.
In this study, we have shown that a functional xylose reductase (XR) from N. crassa can be heterologously expressed in the chloroplast of C. reinhardtii. We assembled and integrated three different expression cassettes containing XR into the chloroplast genome of C. reinhardtii. Protein expression, mRNA levels, and xylitol production were quantitatively compared for each of the transgenic strains expressing these cassettes. Several factors have been identified that affect heterologous protein expression in the chloroplast of C. reinhardtii. The C. reinhardtii chloroplast genome displays a strong codon bias, with A or T (80%) preferred at the third position, and thus codon optimization of the target transgene has been shown to significantly improve protein yields (Franklin et al., 2002; Yoon et al., 2011). This conclusion is supported by our results showing that the C. reinhardtii chloroplasts transformed with optXR accumulated 25-fold more XR protein than native XR gene under the same atpA promoter/5 -UTR. Despite accumulating more XR protein, the strain expressing the optXR gene accumulated 4.3-fold less mRNA than the strain expressing XR. Previous studies have shown that the accumulation of chimeric mRNA in the chloroplast of C. reinhardtii is generally proportional to protein accumulation; however, some notable exceptions have been observed (Barnes et al., 2005), and thus this result appears to be one of these exceptions. Another factor that has been shown to dramatically influence the expression of heterologous proteins in the chloroplast of C. reinhardtii is the selection of the promoter and 5 -UTR used to drive transgene expression. The atpA promoter/5 -UTR was selected for this work based on its ability to produce high levels of chimeric mRNA and recombinant protein in previous studies (Barnes et al., 2005; Noor-Mohammadi et al., 2012). Recently, the fusion of the 16S promoter to the atpA 5 -UTR was shown to boost mRNA and heterologous protein to even higher levels in the chloroplast of C. reinhardtii (Rasala et al., 2011). A similar effect was observed in this work by fusing the 16S promoter to the atpA 5 -UTR to drive optXR expression. The 16S/atpA-optXR strain displayed 5.4-fold higher mRNA levels and 2.9-fold higher protein accumulation than the atpA-optXR strain.
Fig. 5. Comparison of xylitol production from wild-type (WT) C. reinhardtii and the three transgenic strains. No xylitol production was observed in the WT strain. SD of the triplicate measurements is shown in the error bars.
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Although we were able to achieve xylitol production in C. reinhardtii, the 0.05 g/g (xylitol/xylose) yield we obtained using the 16S/atpA-optXR strain was relatively low compared to other xylitol producing microorganisms. For instance, Lee and coworkers achieved a xylitol production yield of 0.92 g/g (xylitol/xylose) by feeding xylose to an engineered strain of C. tropicalis (Lee et al., 2003). Two possible explanations may account for this low yield. First, it is possible that xylose may not be transported efficiently into the chloroplast of C. reinhardtii. The transport of xylose has been shown to be a rate-limiting step for xylose metabolism in other organisms such as the yeast strain S. cerevisiae (Gardonyi et al., 2003). For xylose to be transported into the stroma of the chloroplast of C. reinhardtii, it must pass through multiple membranes including the plasma membrane and the inner and outer membranes of the chloroplast. Passive diffusion through each of these membranes more than likely accounts for the small amount of conversion observed in this study. The outer membrane of C. reinhardtii also does not appear to contain any xylose transporters based on an analysis of its genome sequence. It is possible that the additional expression of xylose transporters in C. reinhardtii would result in higher xylitol yields. Sugar transporters have been successfully introduced into several microalgae strains, including C. reinhardtii, to facilitate the transfer of glucose across the outer membrane and enable heterotrophic growth (Doebbe et al., 2007). Another possible explanation is that there may not be a sufficient amount of the NAD(P)H cofactor available in the chloroplast of C. reinhardtii. The XR from N. crassa uses both NADPH and NADH, thus the depletion of these factors could result in a cofactor imbalance in the chloroplast. Cofactor imbalances have been shown to be a major factor in controlling xylitol accumulation in several yeast strains (van Dijken and Scheffers, 1986; Young et al., 2010). 5. Conclusions In this work, we show that xylitol production in C. reinhardtii is feasible by genetically modifying the chloroplast genome. Despite achieving xylitol production, the overall yield (0.05 g xylitol/g xylose) from C. reinhardtii was relatively low. Further yield improvements are therefore necessary to make the engineered C. reinhardtii strain competitive with other xylitol producing microorganisms. Another possible route for enabling xylitol production in C. reinhardtii would be to target the nuclear genome. Targeting the nuclear genome may alleviate some of the issues involved with transporting xylose into the chloroplast; however, this approach is likely to run into difficulties because transgene silencing remains a significant obstacle to the expression of recombinant proteins in the nucleus of C. reinhardtii (Specht et al., 2010). Acknowledgments This work was supported by NSF EPSCoR award EPS0814361 to the State of Oklahoma and NSF award CHE1048784. A.P. and S.N. were supported by Conoco-Phillips fellowships. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jbiotec. 2013.04.002.
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