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Environmental and Experimental Botany 110 (2015) 19–26

Contents lists available at ScienceDirect

Environmental and Experimental Botany

jo ur nal homep ag e: www.elsev ier .com/ locate /envexpbot

imultaneous overexpression of cyanidase and formateehydrogenase in Arabidopsis thaliana chloroplasts enhanced cyanideetabolism and cyanide tolerance

ashad Kebeisha,1, Mohamed Aboelmyb,c,1, Aymen El-Naggara, Yassin El-Ayoutya,hristoph Peterhanselb,∗

Zagazig University, Faculty of Science, Botany Department, Plant Biotechnology Laboratory, Zagazig, EgyptLeibniz Universitaet Hannover, Institute of Botany, 30419 Hannover, GermanyMinia University, Faculty of Science, Institute of Botany, Minia, Egypt

r t i c l e i n f o

rticle history:eceived 8 July 2014eceived in revised form 29 August 2014ccepted 15 September 2014

eywords:yanideormatehytoremediationrabidopsisynthetic biology

a b s t r a c t

Cyanide is a strong inhibitor of diverse metabolic reactions and easily absorbed by organisms. However,cyanide is also a byproduct of plant and microbial metabolism. This is why these groups of organismscontain pathways for cyanide detoxification. Large amounts of cyanides are also produced by industriesand are today mostly removed by physical and chemical methods. Phytoremediation can provide an alter-native to these techniques, but existing cyanide concentrations at contaminated sites often exceed thecapacities of plant metabolism. In this study, we overexpressed a bacterial cyanidase together with a plantformate dehydrogenase in Arabidopsis thaliana in order to establish a synthetic pathway for cyanide degra-dation. Simultaneous overexpression of both enzymes would ultimately result in the formation of CO2 andNH3 from cyanide. Both enzymes were targeted to chloroplasts and shown to be active in planta. Whenplants were spray-inoculated with cyanide, overexpressors of the synthetic cyanide degradation pathwayshowed less reduction in leaf pigment contents, lower induction of antioxidant enzymes, and reducedgrowth retardation compared to controls. Growth on cyanide was also tested for seedlings germinating

on agar, plants in hydroponics, and plants growing in sand. In all three assays, plants overexpressing thesynthetic pathway for cyanide degradation showed enhanced growth and biomass accumulation com-pared to controls. Gas exchange measurements confirmed enhanced stress resistance of transgenic plantsand suggested that cyanide degradation to CO2 increased the leaf internal CO2 concentration. Results arediscussed in comparison to other approaches for cyanide phytoremediation.

© 2014 Elsevier B.V. All rights reserved.

. Introduction

The cyanide anion (CN−) is a highly toxic compound becauset is easily absorbed by plants and a strong inhibitor ofytochrome c oxidase in the mitochdonrial respiratory electronransport chain (Way, 1984). In addition, other enzymes such asibulose-1,5 bisphosphate carboxylase/oxygenase (Wishnik and

ane, 1969) and enzymes involved in reactive oxygen scavengingKaruppanapandian et al., 2011) are inhibited by cyanide.

Abbreviations: CYND, cyanidase; FDH, formate dehydrogenase.∗ Corresponding author. Tel.: +49 511 7622632.

E-mail address: cp@botanik.uni-hannover.de (C. Peterhansel).1 Both authors contributed equally to this publication.

ttp://dx.doi.org/10.1016/j.envexpbot.2014.09.004098-8472/© 2014 Elsevier B.V. All rights reserved.

Cyanide is a natural metabolite in bacteria, fungi and plants.Many plants including fruit trees and Sorghum produce cyanogenicglucosides that are actively cleaved to produce CN− for defense(Zagrobelny et al., 2004). In addition, cyanide is an equimolarby-product of ethylene biosynthesis originating from the non-catalyzed breakdown of cyanoformate (Peiser et al., 1984) andmight act in a positive feedback loop as an activator of enzymesinvolved in ethylene biosynthesis (Mc Mahon Smith and Arteca,2000).

Plants also possess enzymes for cyanide assimilation. A majorenzyme is beta-cyanoalanine synthase (E.C. 4.4.1.9) that convertscysteine and cyanide to beta-cyanoalanine. Beta-cyanoalanine

might be further converted to asparagine with ammonia beingreleased (Piotrowski and Volmer, 2006). This reaction is cat-alyzed by cyanoalanine hydratase (E.C. 4.2.1.65), a member ofthe superfamily of nitrilases that contains enzymes with different

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unctions related to N-metabolism (Brenner, 2002). Alternatively,eta-cyanoalanine might also act as a signal molecule in develop-ental processes (Garcia et al., 2010) or stress responses (Garcia

t al., 2013; Liang, 2003; Machingura and Ebbs, 2010). Beside beta-yanoalanine synthase, enzymes like rhodaneses or thiosulfateulfurtransferases (EC 2.8.1.1) also participate in cyanide degra-ation in plants by formation of thiocyanides (Papenbrock et al.,011).

Different from plants, bacteria and fungi have hydrolyticnzymes for cyanide degradation. Cyanide hydratases (E.C..2.1.66) are often found in fungi and catalyze the conversion ofyanide to formamide which subsequently decomposes to CO2 andmmonia (Wang et al., 1992). Bacteria mainly contain cyanide dihy-ratases (cyanidases, CYND, E.C. 3.5.5.1) (Ingvorsen et al., 1991;

andhyala et al., 2003). These enzymes catalyze the conversion ofyanide to formate and NH3 (Harris and Knowles, 1983; White et al.,988). Both hydratases and dihydratases belong to the nitrilaseuperfamily. Apart from hydrolytic degradation, microorganismsan also degrade cyanides with monooxygenases (Qian et al., 2011).

Large amounts of cyanides are used by industries such as goldnd silver mining, coal gasification, steel manufacturing, or chem-cal polymer synthesis (Donato et al., 2007; Eisler and Niemeyer,004). These cyanides may accumulate in wastewaters and soils.oday, they are mostly removed by physical and chemical methodsPalmer et al., 1988). An alternative to these methods is biodegra-ation by plants or microorganisms (Baxter and Cummings, 2006;bbs, 2004; Ebbs et al., 2010; Larsen and Trapp, 2006; Trapp et al.,003). However, biological degradation systems might be easilyverloaded with cyanides and this can result in irreversible destruc-ion of the system (Ebbs, 2004).

In order to improve the efficacy of biodegradation mechanisms,ransgenic overexpression of proteins involved in cyanide degra-ation is a useful approach. Recently, it has been shown thatverexpression of a beta-cyanoalanine hydratase in the modellant Arabidopsis thaliana resulted in improved cyanide tolerance

f seedlings grown on agar plates containing cyanide (O‘Learyt al., 2013). The additional enzyme was derived from bacteria, butomologous to the endogenous Arabidopsis protein NIT4. Alter-atively to boosting existing pathways, different mechanisms for

ig. 1. Schematic representation of FDH and CYND gene expression cassettes. The binary constitutive CaM promoter (p35SS) and the 5′ UTR of the Cab22L tobacco leader peptnd formate dehydrogenase (FDH, At5g14780, gi 145358030, E.C. 1.2.1.2) separately andanked by the scaffold attachment region (SAR) of the tobacco RB7 gene (gi3522871) any scaffold attachment region (SAR) of the tobacco RB7 gene (gi3522871) and the 3′ UTR oeparated by a transcriptional blocker sequence (TB) from phage Lambda (gi215104). pcTPuberosum, gi162946536), scTP: a chloroplast targeting peptide of the Rubisco small subu

rimental Botany 110 (2015) 19–26

cyanide degradation might be combined in one organism. Here,we report the overexpression of a cyanidase (CYND) gene from thebacterium Pseudomonas stutzeri in Arabidopsis. The approach wascombined with simultaneous overexpression of the endogenousformate dehydrogenase (FDH) that can convert formate resultingfrom the cyanidase reaction to CO2. This novel synthetic path-way would allow for complete decomposition of cyanide to CO2and NH3, two gases that could be efficiently integrated into plantanabolism (Peterhansel et al., 2010). Cyanide resistance and phys-iology of the plants were investigated in this study.

2. Materials and methods

2.1. Plasmid constructs and plant transformation

The coding sequence for formate dehydrogenase (FDH,gi145358030) in translational fusion to a chloroplast targeting pep-tide was chemically synthesized. NcoI and AscI restriction siteswere added to the 5′ end and 3′ end, respectively. The binaryplant expression vector pTRAK, a derivative of pPAM (gi13508478,see Fig. 1), was opened with NcoI and AscI and the synthesizedfragment was ligated into the vector. The expression cassette forcyanidase (CYND, gi1167514) was completely chemically synthe-sized including the promoter, untranslated regions, the chloroplasttransit peptide, and the transcription blocker (TB) that was usedto separate the CNAD and FDH genes (see Fig. 1). The constructwas flanked by NotI and PstI sites. pTRAK-FDH was opened withNotI and PstI and the cyanidase expression cassette was clonedinto these sites.

Floral dip transformation of Arabidopsis plants was performedas described by Clough and Bent (1998). Briefly, Arabidopsis plantswere grown under short day conditions (8 h light and 16 h darkat 20–22 ◦C) to produce biomass. Afterwards, plants were trans-ferred to long day conditions (16 h light and 8 h dark at 23–25 ◦C)

to induce flowering. Agrobacteria harboring the expression con-structs were grown and flowers of Arabidopsis plants were dippedin the Agrobacterium solution for 3 to 10 min. The dipped plantswere covered over night (16 h) in order to maintain high humidity.

plant expression vector pTRAK (A), a derivative of pPAM (gi13508478) containingide (TL) was used for the expression of cyanidase (CYND, gi1167514, E.C. 3.5.5.1)

together in the chloroplast of A. thaliana plants. The FDH expression cassette (B) isd the 3′ UTR (3′g7, gi307557069). The CYND expression cassette (C) is also flankedf CaM (pA35S). FDH and CYND expression cassettes in the double construct (D) are: a chloroplast targeting peptide of the Rubisco small subunit from potato (Solanumnit from sun flower (Helianthus annus, gi18809).

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he plants were then transferred to long day growth conditionsntil plant maturation and seed harvesting.

.2. Growth assay conditions

Four different systems were used for growth assays underyanide stress. For determination of root growth in seedlings, plantsere grown on vertical Murashige and Skoog (MS) medium agarlates with or without 250 �M KCN (plate assay). For determina-ion of rosette size, fresh weight and dry weight, plants were grownn a hydroponic system in MS medium under short day conditions8 h/16 h, 22/20 ◦C, day/night). After two weeks, plants were treatedith MS supplemented with 50 �M KCN for another two weeks

hydroponic assay). Alternatively, plants were grown in sand andatered with MS medium. After four weeks, plants were treatedith MS supplemented with 300 �M KCN for another two weeks

n 3-day intervals (sand assay). KCN was also applied via the leafurface to six weeks old plants grown in soil. Plants were sprayed for

weeks in 2-day intervals with 2 mM KCN in a closed transparentox (foliar spray assay).

.3. Recombinant protein expression and enzymatic assays

For CYND and FDH assays with recombinant protein overex-ressed in bacteria, pET-CYND and pET-FDH, respectively, wereransformed into the bacterial strain BL21 (Novagen, Darmstadt,ermany). The empty vector pET22b+ was used as a negativeontrol. For protein expression, a 0.5 L culture was grown in LB-edium to an OD600 of 0.4. Protein expression was induced by

ddition of 1 mM IPTG and culture growth was continued for 2 ht 37 ◦C. Cells were washed once in 10 mM potassium phosphatepH 7.5) and resuspended in the same buffer (15–20% (v/v) cell sus-ension). Cells were lysed in bacterial lysis buffer (20 mM Tris–HClH 7.5, 300 mM NaCl, 10% glycerol, 5 mM DTT, 0.1% Triton X-100,

mg/ml lysozyme, and 2 U DNAse I) for 1 h on ice, and the extractas centrifuged for 30 min at 30,000 × g at 4 ◦C. The activity of CYND

n the supernatant was measured following the protocol describedy Watanabe et al. (1998). The activity of FDH in the supernatantas measured following the protocol described by Höpner andnappe (1974). FDH assay buffer contained 57 mM sodium phos-hate buffer (pH 7), 50 mM formate, and 1.1 mM NAD+ in a finalolume of 1 ml. The activity was determined spectrophotometri-ally at 340 nm.

For CYND and FDH assays from plants, chloroplasts were iso-ated from 4 weeks old A. thaliana plants as described by Kebeisht al. (2007). Approximately 5 g leaf material was ground in 50 mlrinding buffer (50 mM HEPES-KOH pH 7.5, 1 mM MgCl2, 1 mMDTA, 1 g l−1 BSA, 0.2 g l−1 sodium ascorbate, 0.3 M mannitol, 5 g l−1

olyvinylpyrrolidone). After filtration through three layers of Mir-cloth, the solution was centrifuged at 1000 × g for 10 min. Theellets were resuspended in 1 ml SH-buffer (50 mM HEPES-KOHH 7.5, 0.33 M sorbitol), and 0.5 ml of this solution was loaded onto

1-ml 35% Percoll gradient (35% Percoll, 65% SH-buffer). The gradi-nt was centrifuged for 5 min at 500 × g. The chloroplast pellet wasashed in 1 ml SH-buffer and chloroplast protein was extracted in

00 �l extraction buffer (50 mM HEPES-NaOH pH 7.5, 2 mM EDTA, mM MgCl2, 0.1% Triton X-100, 20% glycerol). CYND activity waseasured following a protocol modified from Qian et al. (2011).

YND assay buffer contained 50 mM phosphate buffer (pH 7.5)nd 10 mM KCN as substrate. The reaction was started by adding0 �g of chloroplast protein extract and stopped by adding oneolume of Nessler’s reagent (Sigma Aldrich, Germany) at different

ime intervals. The amount of ammonia produced was colorimet-ically measured at 480 nm and compared to a dilution series ofH4Cl standard. Formate dehydrogenase activity was measuredased on the formate dependent oxidation of NAD+ (Höpner and

rimental Botany 110 (2015) 19–26 21

Knappe, 1974). For the determination of antioxidant enzyme activ-ities, 100–150 mg leaves were harvested from 6 to 7 weeks old A.thaliana plants after foliar spray application of 2 mM KCN twice intwo days intervals. Leaf material was collected 24 h after the sec-ond application of KCN, ground in liquid nitrogen to a fine powder,and homogenized in cold phosphate buffer (0.05 M, pH 7.5). Thehomogenate was centrifuged at 12,000 rpm for 20 min at 4 ◦C. Thesupernatant was used for the determination of catalase activity,peroxidase activity, and superoxide dismutase activity (Xu et al.,2008).

2.4. Chlorophyll and carotenoid determination

Pigments were extracted from six weeks old plants after threetimes foliar spray application of 2 mM KCN in three-day intervals.Leaves were collected 24 h after the last application. Chlorophyllcontents were measured as described by Nybom (1955). 100 mgleaves were harvested in a 2 ml reaction tube and immediatelyfrozen in liquid nitrogen. The leaf samples were ground and 1 ml80% acetone was added. Samples were vortexed vigorously andcentrifuged at 30,000 × g for 10 min. Extinctions at 663 nm, 645 nm,and 440 nm were measured and pigment concentrations were cal-culated from these values.

2.5. Ammonia release assay

Ammonia release assay was performed as described by Rasco-Gaunt et al. (1999). Two Arabidopsis leaves were harvested andplaced in a 1.5 ml reaction tube containing 1 ml of incubationmedium (50 mM potassium phosphate pH 5.8, 2% sucrose, 0.1%Tween 20, 0.1 mg l−1 2,4-dichlorophenoxy acetic acid, 25 mg l−1

phosphinotricin, with or without 2 mM KCN). As a blank, two reac-tion tubes containing only incubation medium were used. Thetested samples were incubated under 100 �E light intensity for 16 hat 26 ◦C. 100 �l from each sample were transferred to a new 1.5 mlreaction tube containing 0.5 ml of reagent-I (0.21 M sodium sali-cylate, 0.085 M trisodium citrate, 25 g l−1 sodium tartrate, 0.4 mMsodium nitroprusside). Samples were mixed and 0.5 ml of reagent-II (0.75 M NaOH, 2.3 mM sodium dichloroisocyanurate) was added.Samples were incubated in the dark at 37 ◦C and then room temper-ature for each 15 min. The presence of ammonium ions in the testedsamples resulted in an emerald green to dark blue color whichwas quantified by measuring the extinction at 655 nm comparedto ammonium standards.

2.6. Quantitative RT-PCR

RNA was prepared from A. thaliana leaves following the BCP(1-bromo-3-chlorpropane) protocol (Chomczynski and Mackey,1995). Preparation of first strand cDNA was performed as describedby Niessen et al. (2007). Quantitative PCRs were performed on anABI PRISM 7300 Sequence Detection System (Applied Biosystems,Darmstadt, Germany) in the presence of SYBR Green following themanufacturer’s instructions. Reaction mix was obtained from Invi-trogen, Karlsruhe, Germany, and oligonucleotides were purchasedfrom Metabion, Planegg, Germany. For the detection of FDH trans-cripts, primers 5′-GCT GTT GTT GAT GCT GTT GAA-3′ and 5′-TCCTTA GGA GCT GGT TGT GG-3′ were used. For the detection of CYNDtranscripts, primers 5′-GCC TAA CCC AGT TGT CAG AAA-3′ and 5′-GCA AGG AAC GCA CTA CTG G-3′ were used. For the detection of

Actin2 transcripts, primers 5′- GGT AAC ATT GTG CTC AGT GGTGG-3′ and 5′-GGT GCA ACG ACC TTA ATC TTC AT-3′ were used.The final primer concentration was 200 nM in the reaction mix-ture. Amplification conditions were 10 min of initial denaturation

22 R. Kebeish et al. / Environmental and Experimental Botany 110 (2015) 19–26

Fig. 2. Characterization of FDH and CYND enzymes used in this study. ((A) and (B)) Bacterial overexpression of CYND (A) and FDH (B). Total protein was isolated from bacteriaand tested for activity of the recombinant enzyme. (C) Amount of FDH (black bars) and CYND (gray bars) transcripts relative to Actin2 transcripts in three independentA from

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that CYND can release NH3 from cyanide in planta.

Fig. 3. Ammonia release assay upon exposure to KCN. Two detached plant leavesfrom 6–7 weeks old transgenic A. thaliana plants were incubated in the absence(−KCN) or presence (+KCN) of 2 mM KCN and NH3 release was measured. Azy-gous: Segregants from FDH + CYND plants that lost the transgene, Ev: empty vector

rabidopsis lines overexpressing the FDH + CYND construct. Samples were collectedD) and FDH (E) in chloroplasts isolated from two independent Arabidopsis lines ompty vector construct. Each data point shows the mean of three independent repl

t 95 ◦C, followed by 40 cycles each of 15 s denaturation at 95 ◦Cnd 1 min combined annealing and extension at 60 ◦C.

.7. Gas exchange measurements

Plants grown in sand and watered with MS + 300 �M KCNor two weeks (sand assay, see above) were used for gasxchange measurements using the LI-6400 system (Li-Cor,incoln, NE). Parameters were calculated with the softwareupplied by the manufacturer. Measuring conditions werehoton flux density = 1000 �mol m−2 s−1, chamber tempera-ure = 26 ◦C, flow rate = 100 �mol s−1, relative humidity = 60–70%,nd CO2 = 400 ppm. Plant leaves were allowed to adapt for0–30 min to the measuring chamber before each measurement.

. Results

.1. Establishment and characterization of transgenic lines

We wanted to test whether overexpression of cyanidase (CYND,rom P. stutzeri) could confer enhanced cyanide resistance to plantsnd whether formate resulting from the reaction could be fur-her converted to CO2 by formate dehydrogenase (FDH, from A.haliana). Before transfer to plants, the activity of both enzymes wasested in crude extracts from bacteria overexpressing the respec-ive cDNA (Fig. 2A and B). Significant increases in activity relativeo the empty vector controls could be observed for both enzymes inpecific assays. Thus, both cDNAs encoded the expected enzymaticctivities.

The genes were transferred separately (CYND, FDH) or in com-ination (CYND + FDH) to Arabidopsis plants. The enzymes wereonstitutively expressed, but targeted to chloroplasts to make bestotential use of CO2 resulting from the combined reactions of CYNDnd FDH. Transgenic lines were selected based on kanamycin resis-ance. Transgene expression was initially tested by quantitativeCR in the CNYD + FDH line. As shown in Fig. 2C for three inde-endent lines, both RNAs accumulated in transgenic plants. LinesYND + FDH-4 and CYND + FDH-6 showed higher expression of bothransgenes compared to line number 2. We further tested activ-ty of CYND and FDH in extracts from chloropasts isolated fromYND + FDH plants in order to proof correct localization of thenzymes (Fig. 2D and E). As controls, chloroplast extracts from WT

lants and plants transformed with the empty vector (EV) weresed. A three-fold (CYND) to five-fold (FDH) increase in activityver background was observed in these assays indicating that bothnzymes accumulated in chloroplasts and were functionally active.

the youngest fully expanded leaf 6 h after onset of light. (D) and (E) Activity of CYNDressing the FDH + CYND construct. WT = Wildtyp; EV = plants transformed with an

± SE.

Because NH3 is a product of the CYND reaction, NH3 release fromleaf discs of transgenic plants that were incubated in medium con-taining 2 mM KCN or not (−KCN) was measured. NH3 (re-)fixationin the GS/GOGAT cycle is inhibited in these assays by addition ofphosphinothricin (Rasco-Gaunt et al., 1999). Two controls wereincluded: (1) Azygous plants that were derived from CYND + FDHtransformants, but lost the transgene by segregation, and (2) anempty vector control (EV). As shown in Fig. 3, in the absenceof KCN, NH3 was released from all leaf discs to a similar extent(gray bars). This amount of NH3 is probably derived from glycinedecarboxylation during photorespiration. Addition of KCN (blackbars) did not significantly affect ammonia release in the controllines or the FDH overexpressors. However, a significant two-foldincrease was observed in CYND overexpressors and three indepen-dent CYND + FDH lines overexpressing both enzymes. This indicates

transformants, FDH: plants transgenic for formate dehydrogenase, CYND: plantstransgenic for cyanidase. FDH + CYND: three independent plant lines transgenic forformate dehydrogenase and cyanidase. Data are means from at least three inde-pendent plants for each genotype ± SE. Asterisks represent statistically significantdifferences compared to azygous control plants (** p < 0.01, *** p < 0.001).

R. Kebeish et al. / Environmental and Experimental Botany 110 (2015) 19–26 23

Fig. 4. Pigment accumulation (A), antioxidant enzyme activity (B) and growth ((C)–(E)) after foliar spray inoculation of transgenic Arabidopsis plants with cyanide. Six weeksold plants were sprayed three times in 3-day intervals with 2 mM KCN. Azygous: segregants from FDH + CYND plants that lost the transgene, EV: empty vector transformants,F idasea otypec

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DH: plants transgenic for formate dehydrogenase, CYND: plants transgenic for cyannd cyanidase. Data are means from at least three independent plants for each genontrol plants (* p < 0.05, ** p < 0.01, *** p < 0.001).

.2. Physiological characterization

Initial tests with WT plants suggested that spraying cyaniden Arabidopsis leaves (foliar spray) resulted in a loss of pig-entation. This assay was used to test resistance of transgenic

ines to foliar cyanide application. Fig. 4A shows chlorophyll andarotenoid contents of the transgenic lines in comparison to azy-ous controls after three times application of 2 mM cyanide inhree-day-intervals. Whereas FDH overexpressors did not differrom the control, we observed higher contents of all tested pig-

ents in the CYND overexpressors. The double overexpressorshowed a further increase in chlorophyll a contents compared toingle overexpressors. Thus, KCN-induced loss of pigmentation wasfficiently alleviated by CYND overexpression and double transfor-ants performed even better.Antioxidant enzymes catalase, peroxidase, and superoxide dis-

utase were induced after foliar spray of cyanide. We thereforeeasured the activities of these enzymes in extracts from leaves of

he different plant lines (Fig. 4B). Again, the FDH line did not differ

rom the azygous control whereas CYND and CYND + FDH plantshowed reduced induction of these enzymes (−20% for peroxidasend −80% for both superoxide dismutase and catalase) by KCN.

ig. 5. Growth of transgenic Arabidopsis plants and controls on agar plates (A), in hydroplants that lost the transgene; FDH + CYND-4: plant line transgenic for formate dehydrogt least six plants per genotype ± SE. Asterisks represent statistically significant differenc

. FDH + CYND: three independent plant lines transgenic for formate dehydrogenase ± SE. Asterisks represent statistically significant differences compared to azygous

3.3. Growth assays

We also tested plant growth in the cyanide foliar spray assay.Six weeks old plants were sprayed with 2 mM KCN in two-day-intervals for two weeks. Before the treatment, rosette diameterswere similar for all lines (Fig. 4C). After the treatment, azygousand empty vector controls did not show further growth. How-ever, rosette diameter of transgenic lines increased by 15% (CYND)to 48% (CYND + FDH line 6) during the treatment. At the end ofthe treatment, fresh weight (FW) and dry weight (DW) of testplants was determined (Fig. 4D and E). Single gene (CYND or FDH)overexpressors showed higher FW and DW than control plants. Afurther strong increase to about four-fold higher values comparedto controls was observed for two independent CYND + FDH lines.We concluded that CYND + FDH lines showed best resistance tofoliar spray application of KCN and therefore concentrated on theseplants for further characterization.

Three additional assays for evaluation of cyanide resistancewere used. In all three assays, KCN was supplied via the root system

as would be expected on contaminated soil. Again, a pool of azy-gous plants derived from the same mother plant was used as thecontrol in all assays. First, plants were germinated on vertical agar

onic cultures ((B)–(C)) or in sand ((D)–(E)). Azygous: segregants from FDH + CYNDenase and cyanidase. Data are means of three independent experiments each withes compared to azygous control plants (* p < 0.05, ** p < 0.01, *** p < 0.001).

24 R. Kebeish et al. / Environmental and Expe

Fig. 6. Growth of transgenic Arabidopsis plants and controls on vertical agarpFf

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lates containing 250 �M KCN (+KCN) or not (−KCN). Azygous: segregants fromDH + CYND plants that lost the transgene; FDH + CYND-4: plant line transgenic forormate dehydrogenase and cyanidase.

lates containing KCN and root length was determined (Figs. 5And Fig. 6). Average root length was significantly increased from.6 to 2.5 cm (+56%) in CYND + FDH plants compared to controls.owever, root length was almost identical for both genotypes in

he absence of KCN. Second, plants were grown in hydroponics forwo weeks without KCN and then KCN was added for two weekso half of the plants (Fig. 5B and C). Rosette diameter and FW weresed as indicators for KCN resistance. CYND + FDH plants and con-rols showed very similar growth when KCN was omitted from the

edium. However, KCN addition resulted in a reduction of bothosette diameter (Fig. 5B) and FW (Fig. 5C) by approximately 30%elative to untreated plants in the azygous control line. In contrast,he CYND + FDH line was hardly affected under these conditions+3% for FW and −10% for diameter). As a third assay, we grewlants in sand supplemented with MS medium for four weeks andhen applied MS + KCN for another two weeks. Again, rosette diam-ter and FW were recorded (Fig. 5D and E). No differences werebserved between the genotypes in the −KCN controls. Additionf KCN reduced rosette diameter by 30% and FW by 50% in theontrol. In the CYND + FDH line, diameter and FW were both onlyeduced by 17% when compared to untreated plants. These assaysrovided independent evidence that CYND + FDH double overex-ressors performed superior when KCN was supplied via the rootystem.

.4. Gas exchange characteristics

If cyanide would be completely degraded by CYND and FDH, NH3nd CO2 were the ultimate reaction products. Our previous assaysad shown that NH3 was released from the CYND reaction. In ordero provide evidence that CO2 was formed by FDH, we determinedas exchange characteristics of CYND + FDH and control lines afterwo weeks of KCN treatment. In parallel, plants were grown without

CN treatment and tested (Table 1).

As observed before in growth assays, CYND + FDH plants andontrol plants performed very similar under non-stressed condi-ions (−KCN). KCN treatment reduced CO2 assimilation rate by 14%

able 1as exchange measurements of transgenic lines and azygous controls in the presence and

−KCN

Azygous

Assimilation rate (�mol CO2 m−2 s−1) 15.11 (±0.58)

Intercellular CO2 concentration (Ci , �mol mol−1) 211.31 (±9.2)

Stomatal conductance (mmol H2O m−2 s−1) 0.152 (±0.016)

Transpiration (mmol H2O m−2 s−1) 1.81 (±0.19)

p < 0.05.** p < 0.01.

*** p < 0.001 relative to azygous control.

rimental Botany 110 (2015) 19–26

in the control whereas CYND + FDH plants remained unaffected.Similar trends were observed for transpiration (−14% for control vs.+7% for CYND + FDH) and stomatal conductance (−14% for controlvs. +10% for CYND + FDH). Interestingly, leaf internal CO2 concen-tration was largely unaffected in control plants by the treatment,but significantly increased in CYND + FDH plants by 10%. These datasuggest that leaf internal CO2 might be enhanced in CYND + FDHplants by CO2 release from KCN.

4. Discussion

In this paper, we tested an approach for the enhancementof cyanide resistance in plants. The introduced pathway is syn-thetic, since it combines bacterial and plant elements (CYND andFDH; respectively). Combination of both enzymes would result incomplete oxidation of cyanide to NH3 (resulting from the CYNDreaction) and CO2 (resulting from the FDH reaction). Thus, activ-ity of both enzymes can be estimated in planta by detection oftheir gaseous products. Ammonia release assay (Fig. 3) indicatedthat chloroplast-targeted CYND can convert cyanide to NH3 that isexogenously applied to leaf discs. This was not further enhancedby simultaneous expression of FDH as expected. CO2 release fromformate is more difficult to measure because of the refixationof CO2 in the chloroplast by photosynthesis. However, our gasexchange measurements data indicated that leaf internal CO2 wasenhanced in CYND + FDH plants and that this effect was dependenton KCN application. This suggested that both enzymes cooper-ate in planta in the decomposition of cyanide to two gases thatare both useful substrates for plant biosynthesis. In accordancewith this observation, CYND + FDH plants showed improved growthcompared to CYND plants when cyanide was applied in the foliarspray assay (Fig. 4). However, other stress parameters such aschlorophyll content and antioxidant enzyme activity (Fig. 4) didnot indicate major differences between CYND and CYND + FDHplants.

Photosynthetic measurements indicated that FDH is importantfor CO2 production from cyanide in the leaf. An alternative inter-pretation of the importance of FDH in the synthetic detoxificationpathway would be that FDH simply removed toxic formate result-ing from the CYND reaction that might otherwise accumulate.Formate is a natural endogenous compound in plants resulting fromnon-enzymatic decarboxylation of photorespiratory glyoxylate inperoxisomes (Wingler et al., 1999). It can be used as a C1-donor(Wingler et al., 1999) or can be oxidized by the endogenous formatedehydrogenase that is mainly targeted to mitochondria (Hourton-Cabassa et al., 1998) (although dual targeting to mitochondria andchloroplasts has been reported for Arabidopsis (Herman et al.,2002)). There are different reports about the impact of externallyapplied formate on plants. Whereas low amounts of formate in

the growth substrate might even promote growth (Shiraishi et al.,2000), higher concentrations delay germination or affect plantgrowth (Himanen et al., 2012; Li et al., 2002). Consequently, FDHoverexpression protected plants from formate toxicity (Li et al.,

absence of KCN (means derived from at least six independent plants ± SE).

+KCN

FDH + CYND-4 Azygous FDH + CYND-4

15.39 (±0.35) 13.01 (±0.50) 15.76 (±0.55)**

217.76 (±5.7) 201.41 (±7.9) 237.23 (±4.0)***

0.164 (±0.011) 0.130 (±0.012) 0.149 (±0.013)1.93 (±0.12) 1.56 (±0.15) 1.78 (±0.16)

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002). This detoxification of excess formate might be also the majorunction of FDH in our synthetic cyanide degradation pathway.

Our initial assays with different gene combinations were basedn foliar spray application. However, soil contamination is theajor source of cyanide toxicity to plants (Trapp et al., 2003). Note-orthy, a significant amount of exogenously applied cyanide isot metabolized in roots, but ends up in shoots (Yu et al., 2012).e used three different methods to test resistance of CYND + FDH

lants to cyanide contamination of growth substrates (see Fig. 4).n all these assays, CYND + FDH plants performed significantly bet-er than controls and this effect was always dependent on cyanidepplication indicating that the observed phenotype is not due tony secondary effect of transgene overexpression on growth. Thus,NYD + FDH plants might be suitable for detoxification of soils con-aining increased amounts of cyanides that are not tolerated byther plants. For example, willow trees can detoxify up to 10 mgyanide per kg fresh weight, but rapidly die at concentrations of2 mM (Larsen and Trapp, 2006). Sorghum bicolor can even surviveuch concentrations in the growth substrate by degrading cyanidehat was uptaken (Trapp et al., 2003). Seedlings of the model plantrabidopsis are sensitive to cyanides and show growth defects evenhen exposed to concentrations around 50–100 �M of cyanide

Garcia et al., 2010; Mc Mahon Smith and Arteca, 2000; O‘Learyt al., 2013). In contrast, CYND + FDH plants survived concentra-ions up to 300 �M without visible signs of stress. It remains to behown in further studies whether this approach is also effective inther species that have higher capacities for endogenous cyanideetabolism.It is useful to compare observed effects of CYND + FDH over-

xpression to another recently published transgenic approachowards enhanced cyanide resistance of Arabidopsis that was basedn enhancing the endogenous pathway for cyanide detoxifica-ion (O‘Leary et al., 2013). Plant growth analyses in this paperere focused on seedlings grown on agar plates containing 50 �MCN and are, thus, best compared to our root length data shown

n Fig. 5A (250 �M cyanide in agar plates). In both studies, rootrowth was reduced by approximately 50% after cyanide treat-ent in the WT, but fully restored by the transgenic intervention.hereas carbon and nitrogen from cyanide was assimilated via

ormation of the amino acid asparagine in the endogenous path-ay, the synthetic pathway released gaseous NH3 (Fig. 3) and CO2

Table 1) that were probably re-assimilated by plant primary car-on and nitrogen assimilation pathways. Additional energy andeducing power would be required for re-fixation via gaseous inter-ediates (Peterhansel et al., 2010) compared to direct formation

f amino acids. Together, results indicate that both boosting thendogenous pathway and addition of a synthetic pathway werenstrumental in enhancing cyanide resistance. It would be interest-ng to see whether both approaches can additively enhance cyanide

etabolism in plants and facilitate the use of plants in bioremedi-tion of toxic cyanide contaminations.

cknowledgments

This project was funded by grants from the Egyptian Science andechnology Development Fund (STDF) (GERF 3138) to RK and theerman Ministry of Research (EGY 10/015) to CP. We thank Amirabdel-Motteleb and Moheb Ali for their valuable help with geneloning and technical support.

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