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Autoluminescent Plants

  • Alexander Krichevsky ,

    Affiliation Department of Biochemistry and Cell Biology, State University of New York, Stony Brook, New York, United States of America

  • Benjamin Meyers,

    Affiliation Department of Biochemistry and Cell Biology, State University of New York, Stony Brook, New York, United States of America

  • Alexander Vainstein,

    Affiliation The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, Faculty of Agricultural, Food and Environmental Quality Sciences, The Hebrew University of Jerusalem, Rehovot, Israel

  • Pal Maliga,

    Affiliation Waksman Institute, Rutgers, The State University of New Jersey, Piscataway, New Jersey, United States of America

  • Vitaly Citovsky

    Affiliation Department of Biochemistry and Cell Biology, State University of New York, Stony Brook, New York, United States of America


Prospects of obtaining plants glowing in the dark have captivated the imagination of scientists and layman alike. While light emission has been developed into a useful marker of gene expression, bioluminescence in plants remained dependent on externally supplied substrate. Evolutionary conservation of the prokaryotic gene expression machinery enabled expression of the six genes of the lux operon in chloroplasts yielding plants that are capable of autonomous light emission. This work demonstrates that complex metabolic pathways of prokaryotes can be reconstructed and function in plant chloroplasts and that transplastomic plants can emit light that is visible by naked eye.


Throughout the evolution, bioluminescence has evolved many times and some thirty independent biological light emission systems are still extant [1], in addition to those that may have not survived evolutionary bottlenecks. Among the diverse light-emitting species are bacteria, dinoflagellates, fungi, and insects. The luciferase enzymes, catalyzing the light-emitting reactions in different organisms, show no homology to each other and their substrates, termed luciferins, are unrelated to each other chemically. Luminous bacteria [2], [3] - all gram negative, motile rods - are the most abundant and widely spread of all light-emitting organisms. They are found as free-living species in the ocean, as saprophytes growing on dead marine organisms, as light organ symbionts in fish and squid and other ecological niches. Almost all luminous bacteria, a majority of them being marine species, are classified into three genera: Vibrio, Photobacterium, and Xenorhabdus [3], [4]. Bacterial light-emission enzymatic system, encoded by the lux operon, is highly conserved among various species of luminous bacteria, with the most common architecture of the lux operon represented by luxCDABEG [2], [3]. The bacterial luciferase utilizes flavin mononucleotide and a long-chain aldehyde, derived from fatty lipid biosynthesis pathway, as substrates for the light emission reaction. The luxA and luxB genes encode α and β subunits of the bacterial luciferase, luxC, luxD and luxE encode enzymes involved in the synthesis of aldehyde substrate utilized in the light emission reaction and luxG codes for flavin reductase, which participates in flavin mononucleotide turnover [5], [6].

Until now, expression of various luciferases in plants required exogenous application of luciferins – frequently toxic and high-cost compounds – to achieve only temporary and relatively low light emission levels from live plant tissues [7]. We embarked on an examination to see whether a complete functional bacterial luciferase pathway can be reconstituted in a transplastomic plant to produce both the luciferase and luciferins. Among different independently-evolved luminescent enzymatic apparati, the bacterial light emission system is best suitable for creation of autonomously luminescent plants due to cyanobacterial evolutionary origins of plant plastids [8], [9]. This evolutionary similarity underlies the ability of both plants and bacteria to manufacture riboflavin, from which flavin mononucleotide is produced. The fatty acid biosynthesis pathway, from which the aldehyde substrate is derived, is supported by the same type II fatty acid synthase (FAS II) in both plants and bacteria [10], [11] in contrast to animals and fungi, where fatty acids biosynthesis is mediated by type I fatty acid synthase (FAS I) [12]. Also, plastidal gene expression machinery allows coordinated expression of multitransgene operons [13], [14] and lacks nuclear transgene silencing mechanisms [15], [16], which would be detrimental for expression of a complex pathway involving multiple genes. However, while on the one hand plant chloroplasts share evolutionary origins with prokaryotes, these organelles are not bacteria, and fundamental biological differences between the two exists. For instance, many of the chloroplast encoded open reading frames do not have Shine-Dalgarno sequences, required for bacterial translational initiation, and chloroplast initiation codons are not limited to AUG or GUG as in free living eubacteria [17]. Other differences include coordination of an operon-encoded gene expression, intricacies of transcriptional regulation, posttranscriptional RNA processing, protein folding and quality controls machinery, and other characteristics [18], [19]. It is, therefore, remarkable that higher plants are capable to fully and correctly functionally reproduce a complex enzymatic pathway from an evolutionary unrelated marine organism. While co-expression of a limited number of transgenes in chloroplasts have been investigated [14], a complete and functional foreign biochemical multienzyme pathways have not been reconstructed in plastids. Here, we generated the first truly autonomously luminescent (autoluminescent) transplastomic plants, containing a fully functional bacterial luciferase pathway, which emits visible light detectable by the naked eye.


Generation of Autoluminescent Nicotiana tabacum Plants

We generated two independent lines of Nicotiana tabacum transplastomic plants, carrying the bacterial lux operon from Photobacterium leiognathi. In one line, the lux operon was integrated into the rps12/TrnV locus of the chloroplast genome, whereas, in the other line, it was integrated into a more transcriptionally active TrnI/TrnA locus [20], [21]. To produce these plants, the lux operon, containing the luxCDABEG genes was cloned under the control of the tobacco plastidal Prrn promoter into the plastid transformation vector pCAS3 (see Methods), which carries a spectinomycin resistance selection marker aadA, resulting in pCAS3-aadA-LUX vector. Homologous recombination sites for integration into rps12/TrnV or TrnI/TrnA tobacco plastid genome loci were inserted to flank the aadA-lux expression cassette, resulting in pCA3-LUX-rps12/TrnV or pCA3-LUX-TrnI/TrnA vectors, respectively, and transplastomic plants were generated using standard microbombardment methods and selection on spectinomycin-supplemented media [22], [23].

Once initial shoots had appeared, we used junction PCR to differentiate true transplastomic plants from small ribosomal RNA (rrn16) spontaneous mutants, which are also spectinomycin-resistant [24], [22]. In this approach, one of the PCR primers is located within the expression cassette and the other is positioned within the chloroplast genome, outside of vector sequences, leading to amplification of genome-integrated transgene junction. Fig. 1A illustrates the PCR products predicted to arise from transplastomic plants generated using pCA3-LUX-rps12/TrnV, whereas Fig. 1B demonstrates the results of this junction PCR analysis. The 2.35-kb fragment amplified using primers 78 and 104 and the 2.45-kb fragment amplified using primers 79 and 46 are diagnostic of integration of the entire expression cassette integration into the rps12/TrnV locus (Fig. 1B). Next, we amplified luxB and luxC to further confirm the presence of the lux operon within the transplastomic genome (Fig. 1B). The relatively small (∼155 kb) plastid genome in tobacco is present in thousands of copies per cell [24]. Integration of aadA and the lux operon genes in the LUX-rps12/TrnV genome, and the absence of non-transformed wild-type ptDNA copies has been confirmed by DNA gel blot analyses using plastid targeting sequences, aadA and lux gene probes (Figs. 2 and 3). Transplastomic plants transformed with the pCAS3-LUX-TrnI/TrnA plasmid were identified in a similar fashion, using junction PCR primers specific for TrnI and TrnA, as well as for luxB and luxC. Integration of aadA and the lux operon genes in the LUX-TrnI/TrnA genome, and the absence of non-transformed wild-type ptDNA copies has been confirmed as described for the LUX-rps12/TrnV plants (Figs. 2 and 3). None of the transplastomic plant lines exhibited detectable phenotypic changes in their development or morphogenesis.

Figure 1. PCR analyses to confirm plastid transformation with vector pCAS3-LUX-rps12/TrnV.

(A) Location of PCR primers on the vector and predicted size of junction PCR fragments. Shown are also: the rps12 and trnV plastid genes; aadA, the spectinomycin resistance gene; the lux genes. The drawing is not to scale. (B) Junction PCR fragments obtained by the primers in Fig. 1A using total cellular DNA of a transplastomic plant line as template.

Figure 2. DNA gel blot analysis confirms plastid transformation with vectors pCAS3-LUX-rps12/TrnV (A) and pCAS3-LUX-trnI/trnA (B).

Total cellular DNA isolated from transplastomic leaves was digested with the SmaI restriction endonuclease and probed with a fragment of the vector plastid-targeting region.

Figure 3. DNA gel blot analysis confirms integration of aadA and the lux operon in the LUX-rps12/TrnV (A) LUX-trnI/trnA (B) plastid genomes.

Total cellular DNA isolated from transplastomic leaves was digested with the SmaI restriction endonuclease and probed with a fragment of the vector plastid-targeting region. Shown are also: the rps12 and trnV plastid genes; the trnI and trnA plastid genes; aadA, the selective spectinomycin resistance gene; the lux genes. The drawing is not to scale.

Characterization of Light Emission Properties of the Autoluminescent Plants

Next, we examined the luminescence properties of the LUX-rps12/TrnV and LUX-TrnI/TrnA transplastomic plants. For quantification of light emission, emerging transplastomic or wild-type shoots were placed in scintillation counter vials, incubated in the dark for 5–10 min to eliminate chlorophyll autofluorescence, and photon count was recorded for 20 min. Fig. 4 shows that the transplastomic tissues emitted large quantities of photons of visible light, with LUX-rps12/TrnV and LUX-TrnI/TrnA initially emitting around 3.3×106 and 82.0×106 photons/min (panels A, B, respectively), while background noise was measured using wild type tissue and recorded at only 60–70×103 photons/min. The LUX-TrnI/TrnA plants emitted approx. 25 times more photons from the same amount of tissue than the LUX-rps12/TrnV plants (Fig. 4A, B), presumably due to the higher read-through transcriptional activity of the TrnI/TrnA locus. The decline in the luminescence during the scintillation readings (Fig. 4A, B) most likely resulted from depletion of oxygen in the tightly closed scintillation vials, which is utilized in the light emission reaction.

Figure 4. Quantification and autoradigraphic detection of autoluminescence in transplastomic plants.

(A, B) Scintillation spectroscopy of transplastomic LUX-rps12/TrnV and LUX-TrnI/TrnA tissues (150 mg), respectively, in a Beckman LS 6500 multi-purpose scintillation counter. (C) Photographs (top panel) and autoradiographs of the LUX-TrnI/TrnA plants (bottom panel).

Overnight autoradiography of the LUX-TrnI/TrnA shoots produced a defined and focused image spots around the transplastomic tissue, whereas no such light emission was detected with the wild-type tissue (Fig. 4C). Finally, it was important to demonstrate that the transplastomic plants in fact emit light visible with the naked eye. Indeed, when the fully grown, homoplastomic LUX-TrnI/TrnA plants were placed in a dark room, their glow was clearly seen after about 5–10 min of eye adjustment to darkness. Fig. 5 illustrates the images of these plants as recorded in the dark and in the light, using a standard hand-held consumer camera. No such glow was detected in the wild-type plants.

Figure 5. Visual detection of autoluminescence in LUX-TrnI/TrnA plants.

(A) Photograph taken in the dark with a hand-held consumer camera (Nikon D200; AF-S Micro Nikkor 105.0 mm 1∶2.8 G ED lens; exposures 5 min at f/4.5, 105mm focal length, ISO 3200). (B) Photographs of transplastomic and wild-type plants taken with lights on or off.


In this work, we produced the first example of autoluminescent plants, with the glow clearly visible to the human eye. The basic biochemical machinery required by the lux operon-encoded enzymatic pathway for light emission is very similar across all plant kingdom, making our approach applicable to essentially any plant species. The plant autoluminescence might be further modified in regard to its intensity through the use of different promoters, modification of luciferase substrate levels in the cell, or enhancement of the luciferase catalytic activity via directed evolution [25]. The color of the emitted light can also be modified via luciferase mutagenesis [26], and glow in specific plant organs can be achieved through the use of chloroplast-targeted nuclear-encoded transcription factors expressed from tissue-specific promoters. Thus, our findings not only enhance our understanding of the fundamental biological processes and expression of complex and fully functional multienzymatic pathways in plant plastids, but also are useful for floriculture industry, particularly since plastid DNA is maternally inherited in most flowering plant species [27], substantially reducing the risks of transgene escape into the environment.

Materials and Methods

Chloroplast transformation vectors

Chloroplast transformation vectors of the pCAS series were constructed using the backbone of the pSAT4-MCS vector (GenBank accession number DQ005466.1; [28]). The CaMV 35S promoter of pSAT4-MCS was replaced by a chloroplast Prrn promoter, cloned as a AgeI/NcoI PCR fragment amplified using forward 5′-TCACCGGTCGCCGTCGTTCAATGAGAATGG-3′ and reverse 5′-GAGCGAACTCCGGGCGAATATCCATGGTT-3′ primers and Nicotiana tabacum plastid genomic DNA as a template, resulting in pCAS3 vector. Then, a spectinomycin resistance gene aadA fused to an rbcL leader sequence was cloned into pCAS3 as a BglII/NcoI PCR fragment amplified using forward 5′-AACCATGGAGTTGTAGGGAGGGATTTATGGGGGAAGCGGTGATCGCC-3′ and reverse 5′-TGGAGATCTTTATTTGCCGACTACCTTGGTGATC-3′ primers and pPZP-RCS2 [28] as a template, producing pCAS3-aadA. The lux operon from Photobacterium leiognathi (GenBank accession number M63594), comprising luxCDABEG, was cloned as an EcoRI PCR fragment amplified using forward 5′-ACAGAATTCCCAAAGGAGATTACATGATTAAG-3′ and reverse 5′- TTGGAATTCTTACGTATAGCTAAATGCATCAG-3′ primers and Photobacterium leiognathi genomic DNA as a template into the same site of pCAS3-aadA, resulting in pCAS3-aadA-LUX.

To allow integration into the rps12/TrnV or TrnI/TrnA loci, the corresponding homologous recombination (HR) sequences were amplified from the Nicotiana tabacum plastid genomic DNA and inserted to flank the lux operon expression cassette in pCAS3-aadA-LUX. For pCA3-LUX-rps12/TrnV, the rps12 HR sequence was first cloned as an AgeI PCR fragment amplified using forward 5′-AGTTAGAACCGGTGAAGTGCTTCGAATCATTGCTATTTG-3′ and reverse 5′-CGATCTAACCGGTTTATCAACTGCCCCTATCGGAAATAGG-3′ primers. The TrnV HR sequence was then cloned into the resulting vector as an NcoI PCR fragment amplified using forward 5′-ATAATGCGGCCGCCAATTGAATCCGATTTTGACCATTATTTTC-3′ and reverse 5′-ATTATGCGGCCGCGTGAAGCAGTGTCAAACCAAAATACC-3′ primers. For pCA3-LUX-TrnI/TrnA, the TrnI HR was first cloned as an AgeI PCR fragment amplified using forward 5′-AGTTAGAACCGGTCTTCGGGAACGCGGACACAGGTGG-3′ and reverse 5′-CGATCTAACCGGTAGATGCTTCTTCTATTCTTTTCCCTG-3′primers. The TrnA HR sequence was then cloned into the resulting vector as a NotI PCR fragment amplified using forward 5′-CTATTATGCGGCCGCACTACTTCATGCATGCTCCACTTGG-3′ and reverse 5′- GAATGATGCGGCCGCCCTATGAAGACTCGCTTTCGCTACG-3′ primers. All constructs were verified by DNA sequencing.

Transplastomic plants

Transplastomic Nicotiana tabacum (cv. Petit Havana) plants were produced by standard chloroplast transformation protocols [22], [23]. Homoplastomy was confirmed by the Southern blot analysis.

Junction PCR primers

Transplastomic LUX-rps12/TrnV plants were identified using primers 78 (5′-TTGAGTATCCGTTTCCCTCC-3′) and 104 (5′-CCAGCAAATCAATATCACTGTGTGG-3′), which produced a 2.35-kb fragment, and primers 46 (5′-CAGATTTATCTGACTTTGATATCTATG-3′) and 79 (5′-AAGCTCATGAGCTTGGTCTTAC-3′), which produced a 2.45-kb fragment. Transplastomic LUX-TrnI/TrnA plants were identified using primers (5′-CGTTCGCAAGAATGAAACTCAAAGG-3′) and (5′-CAACATCACTTTGGGTGATGATAGG-3′) producing approx. 2.8kb fragment and specific for TrnI junction, and primers (5′-CAGATTTATCTGACTTTGATATCTATG-3′) and (5′-CGCTGATTCTTCAACATCAGTCG-3′) producing approx. 2.2kb fragment and specific for the TrnA junction. The luxB and luxC genes were amplified using primer pairs 5′-ATGAATTTCGGGTTATTTTTCC-3′/5′-TTATTTAATAAGGTTATCTTTG-3′ and 5′-ATGATTAAGAAGATCCCAATGA-3′/5′-CTACGGTACAAATACGAGGAAC-3′, respectively. Primers 73 (5′-AATTGAATCCGATTTTGACCATTATTTTC-3′) and 79 (5′-AAGCTCATGAGCTTGGTCTTAC-3′) amplified an intact region of the chloroplast genome for positive controls.

Author Contributions

Conceived and designed the experiments: AK. Performed the experiments: BM. Analyzed the data: AK. Wrote the paper: AK VA PM VC.


  1. 1. Wilson T, Hastings JW (1998) Bioluminescence. Annu Rev Cell Dev Biol 14: 197–230.
  2. 2. Meighen EA (1993) Bacterial bioluminescence: organization, regulation, and application of the lux genes. FASEB J 7: 1016–1022.
  3. 3. Meighen EA (1991) Molecular biology of bacterial luminescence. Microbiol Rev 55: 123–142.
  4. 4. Baumann P, Baumann L, Woolkalis M, Bang S (1983) Evolutionary relationships in Vibrio and Photobacterium. A basis for a natural classification. Ann Rev Microbiol 37: 369–398.
  5. 5. Lin JW, Chao YF, Weng SF (1998) Characteristic analysis of the luxG gene encoding the probable flavin reductase that resides in the lux operon of Photobacterium leiognathi. BBRC 246: 446–452.
  6. 6. Nijvipakul S, Wongratana J, Suadee C, Entsch B, Ballou DP, et al. (2008) LuxG is a functioning flavin reductase for bacterial luminescence. J Bacteriol 190: 1531–1538.
  7. 7. Greer LF III, Szalay AA (2002) Imaging of light emission from the expression of luciferases in living cells and organisms: a review. Luminesc 17:
  8. 8. Reyes-Prieto A, Weber AP, Bhattacharya D (2007) The origin and establishment of the plastid in algae and plants. Annu Rev Genet 41: 147–168.
  9. 9. Gould SB, Waller RF, McFadden GI (2008) Plastid evolution. Annu Rev Plant Biol 59: 491–517.
  10. 10. Slabas AR, Fawcett T (1992) The biochemistry and molecular biology of plant lipid biosynthesis. Plant Mol Biol 19: 169–191.
  11. 11. Rock CO, Jackowski S (2002) Forty years of bacterial fatty acid synthesis. BBRC 292: 1155–1166.
  12. 12. Smith S (1994) The animal fatty acid synthase: one gene, one polypeptide, seven enzymes. FASEB 8: 1248–1259.
  13. 13. Maliga P (2003) Progress towards commercialization of plastid transformation technology. Trends Biotechnol 21: 20–28.
  14. 14. Arai Y, Shikanai T, Doi Y, Yoshida S, Yamaguchi I, et al. (2004) Production of polyhydroxybutyrate by polycistronic expression of bacterial genes in tobacco plastid. Plant Cell Physiol 45: 1176–1184.
  15. 15. Mathieu O, Bender J (2004) RNA-directed DNA methylation. J Cell Sci 117: 4881–4888.
  16. 16. Chen X (2009) Small RNAs and their roles in plant development. Annu Rev Cell Dev Biol 25: 21–44.
  17. 17. Zerges W (2000) Translation in chloroplasts. Biochimie 82: 583–601.
  18. 18. Bock R (2007) Topics in Current Genetics, Cell and Molecular Biology of Plastids. Springer-Verlag Berlin Heidelberg 19.
  19. 19. Sakamoto W (2006) Protein degradation machineries in plastids. Annu Rev Plant Biol 57: 599–621.
  20. 20. Lutz KA, Azhagiri AK, Tungsuchat-Huang T, Maliga P (2007) A guide to choosing vectors for transformation of the plastid genome of higher plants. Plant Physiol 145: 1201–1210.
  21. 21. Verma D, Daniell H (2007) Chloroplast vector systems for biotechnology applications. Plant Physiol 145: 1129–1143.
  22. 22. Lutz KA, Svab Z, Maliga P (2006) Construction of marker-free transplastomic tobacco using the Cre-loxP site-specific recombination system. Nat Protoc 1: 900–910.
  23. 23. Verma D, Samson NP, Koya V, Daniell H (2008) A protocol for expression of foreign genes in chloroplasts. Nat Protoc 3: 739–758.
  24. 24. Svab Z, Maliga P (1991) Mutation proximal to the tRNA binding region of the Nicotiana plastid 16S rRNA confers resistance to spectinomycin. Mol Gen Genet 228: 316–319.
  25. 25. Turner NJ (2009) Directed evolution drives the next generation of biocatalysts. Nat Chem Biol 5: 567–573.
  26. 26. Shapiro E, Lu C, Baneyx F (2005) A set of multicolored Photinus pyralis luciferase mutants for in vivo bioluminescence applications. Protein Eng Des Sel 18: 581–587.
  27. 27. Hagemann R (2004) The sexual inheritance of plant organelles. In: Daniell H, Chase CD, editors. Dordrecht, The Netherlands: Molecular Biology and Biotechnology of Plant Organelles Springer. pp. 93–113.
  28. 28. Tzfira T, Tian GW, Lacroix B, Vyas S, Li J, et al. (2005) pSAT vectors: a modular series of plasmids for autofluorescent protein tagging and expression of multiple genes in plants. Plant Mol Biol 57: 503–516.