Trichoderma atroviride PHR1, a Fungal Photolyase Responsible for DNA Repair, Autoregulates Its Own Photoinduction

Transgenic fungal strains and nucleic acid and protein methods.To obtain null strains, a disruption construct was made as follows. The 5′-flank (2 kb) and 3′-flank (1.73 kb) fragments of phr1 were generated by PCR, using pT3T7Thphr1 5.8 as a template. The 5′ flank, with PstI/NdeI sites, was obtained using primers PL5 and PL6. The 3′ flank, with BamHI/HindII sites, was obtained with primers PL9 and PL10. The green fluorescent protein (GFP) coding region (0.8 kb), with NdeI/SalI sites, was amplified using synthetic GFP as a template, with primers PL7 and PL8. A 1.4-kb fragment containing the hph gene was obtained from either pCSN44 or pUCATPH by digestion with the SalI/BamHI enzymes. All of these fragments were introduced by one-step ligation into a pBlueScript SK vector at PstI/HindIII sites, and the construct was named pC16. The primer sequences were as follows: PL5, 5′GCA ACT GCA GCG CTT CTT GTC ACT TTC T3′; PL6, 5′GTA GAT CCA TAT GTT ACA CAC GGT GGA TTC TTA AAT GAT AAT3′; PL7, 5′GTA GAT CCA TCA TGG TGA GCA AGG GCG AGG AG3′; PL8, 5′ACG CGT CGA CGC AAG ACC GGC AAC AGG ATT CAA3′; PL9, 5′AGC TGG ATC CAT TGG ATA TCG AAA ACG GTT CAT C3′; and PL10, 5′GAT CAA GCT TGT CGG AAG AGA TCA CCA GTG TTT T3′. The construct was transformed as either circular plasmid DNA or linear DNA amplified from the plasmid by PCR with primers PL5 and PL10, by particle bombardment, or by the polyethylene glycol-Ca²⁺ method. MC strains were obtained by cotransformation of spores of T. atroviride IMI 206040 with the plasmids pHatα (8.9 kb) and pT3T7phr1 5.8 (8.5 kb) by particle bombardment as described previously (36). The bombarded plates were overlaid with 400 μg/ml hygromycin B in agar. Transformed strains selected on hygromycin at 200 μg/ml were subjected to monosporic subculture. Null strain candidates were analyzed by PCR for the presence of phr1, using primers PL11 (5′ TGC AAG CGG GAC AAT CAG TTC3′) and PL12 (5′ AAT CCC CAG CCA CCG TTG TTG3′). To detect the presence of gfp, we used primers PL7 and PL8 (see above), and for hph, we used primers HygF (5′ GAG GGC GAA GAA TCT CGT GC3′) and HygR (5′ CAC TGA CGG TGT CGT CCA TC3′). For Southern analysis, 5 μg of total DNA was digested with SalI, resolved in a 0.8% agarose gel, and blotted onto a nylon membrane. For Northern analysis, total RNA was isolated using TriReagent (MRC) following the manufacturer's protocol. Thirty micrograms of RNA was separated by electrophoresis on denaturing formaldehyde gels, blotted onto Hybond N+ membranes (Amersham), hybridized in 7% sodium dodecyl sulfate (SDS), 250 mM phosphate, pH 7, and washed twice for 20 min each time in 5% SDS, 20 mM phosphate, followed by a 5- to 10-min wash with 1% SDS, 20 mM phosphate. Signals were measured using a phosphorimager (Fuji) and Tina v.2.1 software and normalized to total RNA detected by hybridization with an rRNA genomic fragment or by quantitation of ethidium bromide fluorescence on stained gels prior to transfer. In some earlier experiments, signals were quantified by scanning autoradiograms and integration using NIH Image 1.63 software.

To obtain total cell protein extracts, mycelium was lyophilized, ground to a powder, and extracted in Laemmli sample buffer with 1 mM phenylmethylsulfonyl fluoride by vortex mixing with acid-washed glass beads. The extract was heated for 5 min to 100°C and centrifuged for 5 min at 18,000 × g, and 160 μg/lane was separated by SDS-polyacrylamide gel electrophoresis, blotted onto a nitrocellulose membrane, and probed with antiserum to Neurospora photolyase at a 1:500 dilution; the second antibody was goat anti-rabbit (GIBCO) diluted 1:5,000, and detection was done by ECL (Amersham).

Real-time PCR.Approximately 1 μg total RNA was treated with 1 Kunitz unit of RNase-free DNase (QIAGEN) for 20 min at 37°C in New England Biolabs restriction enzyme buffer 4 (total volume, 10 μl), followed by 15 min at 65°C to inactivate the DNase. Four microliters of this reaction mix was reverse transcribed (ImProm-II; Promega) according to the recommended protocol, using random primers; the reaction was done for 1 h at 42°C with 3.75 mM MgCl₂, followed by 15 min at 70°C to inactivate the reverse transcriptase. Four microliters of a 100-fold dilution of each cDNA (in the linear range for all primer pairs, as tested in preliminary experiments on a pooled cDNA sample) was amplified in a 20-μl reaction mix with SYBR green ROX mix (ABgene, Surrey, United Kingdom) and 1 pmol of each primer. Primer pairs (forward and reverse [5′-3′]) were as follows: CGC CTG GCG AGA TTT TTA CA and ACA GAC ATA TGG CCA GTT AAC CAA A for photolyase, GAG CTG AAG GGC ATC GAC TT and CTT GTG CCC CAG GAT GTT G for GFP, and GAT GGG TGT CAA CCA CAA GGA and GCA AGA GGC GTT GGA GAG AAC for glyceraldehyde-3-phosphate dehydrogenase. Reactions were run in duplicate or triplicate in an Applied Biosystems 7000 cycler. Data were analyzed using the $$mathtex$$\(2^{{-}{\Delta}{\Delta}C_{T}}\)$$mathtex$$method (35), with the glyceraldehyde-3-phosphate dehydrogenase primers as the standard primer pair and the maximum fluence as the standard condition. The cycle threshold (C_T) values for the two or three replicate PCRs with each RNA sample were averaged, and then $$mathtex$$\(2^{{-}{\Delta}{\Delta}C_{T}}\)$$mathtex$$values were calculated, followed by averaging of the independent experiments.

Growth and photoinduction conditions.Standard culture conditions were PDYC medium (24 g/liter potato dextrose broth, 2 g/liter yeast extract, 1.2 g/liter casein hydrolysate; all from Difco) at 25°C. For photoinduction experiments, cultures were grown in liquid PDYC medium as described previously (5) for 36 h in total darkness, after which they were exposed to a pulse of blue light of 540 μmol m² or to the light and irradiance required. Samples were kept in the dark and collected 15 min after exposure under a red safelight. Light of different wavelength bands was obtained by filtering the output of cool-white fluorescent tubes or a slide projector (equipped with heat filter 115 [Schott]) through the following filters: B, blue Lee filter 183 or a blue acrylic filter (4); V, violet Lee filter; BG, blue-green Lee filter 124; R, red Lee filter 106 or red dichroic filter (OCLI, Santa Rosa, CA); and G, cyan dichroic filter (OCLI) combined with cutoff filter OG 515 (Schott). For UVA, light from a long-wavelength UV lamp (UVP) passed through a wide-band UG11 filter (Corning). Fluence rates were adjusted with neutral-density filters and measured with a LiCor (Lincoln, NB) quantum photometer (visible region) or UV meter (UVP) for UVA. Fluences used were 78, 420, and 1,050 μmol m⁻².

Complementation of Escherichia coli deficient in photolyase.The coding sequence of phr1 was obtained in two steps, using the genomic clone pT3T7phr15.8 (5) as a template for PCR amplification using Pfu DNA polymerase (Promega). First, two fragments were obtained using the following oligonucleotide pairs, where F indicates sense and R indicates antisense primers: F, 5′CAG GAA TTC GAT GCT CGC GAG GAG CG3′; 2R, 5′GTT CAT ACA GAC ATA TGG CCA G3′; 3F, 5′GGC CAT ATG TCT GTA TGA ACA AGC C3′; and R, 5′GTT CGG GCC CTA GAT CCC TCT CTC CAG CC3′. These two products were combined and used as a template for amplification using primers F and R (including the start and stop codons, whose locations are indicated in bold), resulting in removal of the single intron in phr1. The 1.9-kb final amplified product was digested with EcoRI and ApaI and ligated into pBluescript SK (Stratagene), and the construct was verified by sequencing both strands. This plasmid was used to transform electrocompetent cells of E. coli SY2 (JM107 Δphr::Cm^r ΔuvrA::Km^r ΔrecA::Tet^r) to ampicillin resistance, and the resulting strain was tested for photorepair of UV damage. For photorepair assays, bacterial colonies were isolated from LB plates with 50 μg/ml ampicillin, 50 μg/ml chloramphenicol, 25 μg/ml kanamycin, and 20 μg/ml tetracycline; overnight cultures were plated at different dilutions on LB with 50 μg/ml ampicillin, dried for 30 min in a laminar flow hood, and exposed to the indicated UVC fluences from a cross-linker (Hoefer). The UVC fluence rate was reduced with a neutral filter consisting of several layers of nylon mesh and calibrated with a UVP UV meter.

UVA and blue light induce phr1 expression.We reported previously that the blue region of the spectrum was effective in the induction of phr1 expression (4, 5). In order to compare phr1 induction with the other blue light responses, photorepair and photoconidiation, we mapped the wavelength dependence of phr1 mRNA induction. We used wide-band illumination in defined spectral regions from 300 to 760 nm. Light between 300 and 550 nm is effective for phr1 photoinduction. UVA is the most effective region of the spectrum for phr1 photoinduction, being almost twice as effective as the blue (B) region, followed by the violet (V) and the blue-green (BG) regions. phr1 photoinduction is blind to red light (R) (Fig. 1).

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FIG. 1.

Wavelength dependence of phr1 induction. Northern analysis was performed from induced cultures by fluences of different spectral regions. Data from different blue filters were consistent within the variability and were combined. The peak transmission wavelengths, in nm, of the filters were as follows: UVA, 360 (300 to 390); violet, 412 (330 to 530); blue, 440 (395 to 500) and 458 (380 to 560); blue-green, 505 (450 to 560); green, 505 to 600; and red, 585 to 760 (the numbers in parentheses indicate the ranges for >10% transmission). Relative effectiveness was calculated from densitometric measurements of phr1 and normalized against the 28S rRNA signal, in the linear range (0 to 0.6 of maximum) (see Fig. 7). Bars are means with standard errors (SE) for two to six replicates. The photoreactivation action spectrum is shown schematically as a line for comparison.

Deletion of phr1 results in a loss of photoreactivation, while phr1 overexpression increases photoreactivation capacity.The 62% identity of the deduced amino acid sequence of PHR1 to Neurospora crassa photolyase suggested that phr1 encodes a functional photolyase belonging to class I, type I (5). We tested whether phr1 could complement a CPD photolyase mutant of E. coli, i.e., strain SY2. Cells harboring phr1 had a UVC survival rate of 3 to 4 orders of magnitude higher than that of cells carrying a control plasmid after exposure to photoreactivating light (Fig. 2), indicating that phr1 indeed confers photorepair activity.

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FIG. 2.

Trichoderma phr1 encodes a functional photolyase. E. coli lacking CPD photolyase was complemented by phr1. Circles, E. coli SY2 carrying the phr1 coding sequence (Phr1); diamonds, E. coli SY2 carrying phr1 cloned in reverse orientation (control). Photoreactivation (L; empty symbols) was induced with white light for 30 min at 30 μmol m⁻² s⁻¹; dark (D; filled symbols) samples were incubated in total darkness following UVC exposure. Each point is the mean of two replicates.

To answer the question of whether photolyase plays another role in Trichoderma in addition to its DNA repair function, we generated strains in which phr1 was not present or was overexpressed. Strains lacking the photolyase gene were obtained by gene replacement, using the following approach. Fragments flanking the coding region of phr1 were fused to the hygromycin phosphotransferase gene (hph) as a selectable marker (Fig. 3A). This construct was introduced into Trichoderma, and candidates were screened by PCR. Two transformants, 3.20 and 3.3, showed double homologous recombination of the construct (phr1 is absent and hph is present), while 3.21 showed ectopic integration (Fig. 3B). phr1 MC strains were generated by the introduction of plasmid pT3T7phr1 5.8 (5), containing the full-length genomic clone (Fig. 3C). Multiple integration of the plasmid was confirmed by Southern analysis. Strains carrying one (MC8) and two (MC6 and MC7) extra copies of phr1 were obtained (Fig. 3D).

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FIG. 3.

Construction of transgenic strains. (A) Strategy for replacement of phr1. The construct contains the upstream region of the phr1 coding sequence (5′ flank) ligated to the gfp coding region, followed by the hph gene under the control of the trpC promoter and, finally, by the downstream region of phr1 (3′ flank). The arrow represents the coding region. (B) Identification of phr1-deleted strains (Δphr1) by PCR analysis of transformants. Amplification of phr1 and markers was assayed with genomic DNAs from the indicated strains. 3.20 and 3.3, Δphr1 strains; 3.21, ectopic integration strain. (C) Plasmid pT3T7phr1 5.8 map used for obtaining MC strains. Relevant restriction enzyme sites are shown. Black, plasmid region; gray, phr1 clone. (D) Identification of overexpresser strains (MC) by Southern analysis of the phr1 transformants. A blot of SalI digests of genomic DNAs from the indicated strains was probed with the 5.8-kb phr1 PstI fragment. The 4.8-kb bands correspond to the WT gene, and 3.6-kb bands correspond to the exogenous copy. The 1.2-kb fragment is present in both digests. The densitometric ratio of the 3.6-kb to 4.8-kb bands shows that strains MC6 and MC7 have two extra copies and strain MC8 has one extra copy. V contains the cotransformation vector. (E) PHR1 levels in transgenic strains were determined by Western blotting (top). Ponceau staining is shown as a loading control (bottom). Dark, samples grown in total darkness; light, samples were frozen 30 min after a saturating exposure to blue light (540 μmol m⁻²).

In order to determine whether more phr1 copies indeed correlated with more PHR1 protein, we analyzed the mRNA and protein levels of phr1 in MC strains. We reported previously that, in Trichoderma, low fluences of blue light rapidly induce the expression of phr1 in dark-grown mycelium and conidiophores (5). Consequently, we analyzed whether the phr1 mRNA and protein levels were increased in MC strains in the dark and under photoinduced conditions. Dark levels of phr1 mRNA were not detectable in any of the strains, while under photoinduced conditions phr1 expression was three- to fivefold higher in MC strains than in the WT (data not shown). Western blot analysis using antiserum against the Neurospora photolyase and using dark-grown and photoinduced cultures showed that PHR1 was almost undetectable in the dark but that a polypeptide of the expected size was clearly detectable in photoinduced cultures. This signal was seven- to ninefold more abundant in the lines with two extra copies, whereas no protein could be detected in the Δphr1 strains 3.20 and 3.3 (Fig. 3E).

Δphr1 strains 3.20 and 3.3 were assayed for photoreactivation, by reversion of germination inhibition of UVC-inactivated spores (41), at different UVC doses. UVC-treated Δphr1 spores were unable to restore germination after white light treatment, while WT spores did (Fig. 4A). As expected, Δphr1 strains completely lost photoreactivation ability (Fig. 4B). In contrast, we did not observe an increase in UVC survival rates for PHR1-overexpressing strains compared to that for the WT (data not shown), but a significant increase in their photoreactivation capacity was detected when either dark- or light-grown spores were assayed under nonsaturating, photoreactivating light conditions (Student t test; P < 0.05) (Fig. 4C). Other than the loss of photoreactivation, the loss of phr1 in Trichoderma, as in Neurospora (45), conferred no obvious phenotypes: growth, sporulation, and pigmentation were normal (data not shown).

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FIG. 4.

Photoreactivation of spores from transgenic lines. (A) Loss of photoreactivation in Δphr1 mutant line. One-week-old spores from the WT and a Δphr1 mutant grown in constant light at 30 μmol m⁻² s⁻¹ were either light treated or not, kept in darkness for 16 h, and examined for germination. The percentage of spore germination represents the percentage of photoreactivation. Control, no light treatment; UV, after inactivation by 100 J m⁻² UVC; UV + L, exposed for 30 min to 40 to 60 μmol m⁻² white light irradiation immediately after UVC treatment. (B) Photoreactivation curves from Δphr1 3.3 and 3.20 mutant lines. Spores were exposed to the indicated amounts of UVC (x axis) and then either incubated in the dark (D) or exposed to photoreactivating light treatment (L). (C) Photoreactivation of MC lines. Seven-day-old spores from either cultures grown with a 16-h photoperiod at 30 μmol m⁻² s^−l irradiance or dark-grown spores from MC strains (MC6, MC7, and MC8) and controls (WT and V) were assayed for photoreactivation as described previously (41), except that the photoreactivating light exposure was 15 min from a cool-white fluorescent tube at 40 to 60 μmol m⁻² s⁻¹. Bars represent means ± SE (n = 6). Values are as follows: WT, 23.41 ± 6.17; V, 29.73 ± 7.97; MC6, 47.82 ± 8.32; MC7, 49.72 ± 7.51; and MC8, 59.46 ± 10.65.

Light responsiveness in Δphr1 mutants and phr1 MC strains.If PHR1, in addition to its CPD photolyase function, has any photosensory role, phr1 deletion or overexpression could modify Trichoderma light responsiveness. Therefore, we tested phr1 photoinduction in phr1 null and MC strains. For this purpose, in Δphr1 strains, the phr1 coding region was replaced with the GFP coding region (Fig. 3A). GFP was inserted immediately after the 2-kb left flank of the construct, which is the region ending at the predicted translation start codon (5). GFP photoinduction in the Δphr1 strains 3.20 and 3.3 shows that upon replacement, the upstream region is still able to confer light regulation on the reporter gene (Fig. 5A). PHR1, therefore, is not required for its own induction. In an ectopic integrant, the photoinduction of the resident and transgenic genes was tightly correlated (Fig. 5B), indicating that all the regulatory elements are contained within the 2-kb upstream region represented by the left flank of the construct. Time courses for GFP light induction in Δphr1 strains and for phr1 photoinduction in MC strains were similar in general to that for the WT. In the decay phase, however, the time course was altered (Fig. 5C). While in the WT and overexpression strains the expression levels at 120 min decreased to <25% of the peak, in Δphr1 strains the expression levels were still 65 to 66% of the maximum. The deletion strains may have slow decay or may have lost adaptation, or the stability of the gfp mRNA may be greater than that of the endogenous transcript.

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FIG. 5.

gfp under control of the phr1 promoter is photoinduced in phr1 null strains. (A) Northern analysis was performed on total RNA from dark-grown (D) or light-induced cultures (L) exposed to a fluence of 540 μmol m⁻² blue light. 3.3 and 3.20, Δphr1 mutants, 3.21, ectopic line. (B) Correlation of gfp and phr1 expression in a cell line carrying both the transgene and the resident phr1 copy. Each point represents the gfp signal (x axis) and phr1 signal (y axis) of a single RNA sample. (C) Time course of photoinduction in transgenic lines. Northern analysis was performed on total RNA from dark-grown (0) or light-induced cultures exposed to a fluence of 540 μmol m⁻² blue light. Samples were collected 0, 5, 15, 30, 60, and 120 min after the pulse. The phr1 signal value was normalized against the 28S signal. The curves of phr1 induction for the WT and the transgenic lines are shown. MC6 and MC7 are MC strains, and 3.3 and 3.20 are null mutants.

The light-regulated GFP was used to test whether PHR1 makes any contribution to its own induction. A comparison of photoinduction fluence response curves for the gfp transcript in Δphr1 strains and for phr1 in WT strains shows that the light requirement for half-saturation (k_1/2) of Δphr1 strains was shifted to lower values than those for either WT or ectopic strains (Fig. 6B; Table 1). For individual strains, k_1/2 values were 13.6 μmol m⁻² for strain 3.20 and 19.3 μmol m⁻² for strain 3.3, compared to 32.1 μmol m⁻² for GFP transcripts in the ectopic strain 3.21 and 82.4 μmol m⁻² for phr1 transcripts in the WT. The phr1 promoter light sensitivity was increased fivefold in the Δphr1 strains compared to that of the WT.

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FIG. 6.

Light sensitivity for induction of gfp under control of the phr1 promoter in Δphr1 strains. (A) Northern blot representative of the data used to construct the graph in panel B. The fluence of the light pulse is indicated for each lane. (B) Fluence response curves. Each RNA sample was from one or two mycelial colonies; data are from phosphorimager scanning of the RNA blot hybridizations. The phr1 and gfp signal values were normalized against the rRNA signal, and the data were then normalized for overall changes in hybridization intensity between experiments by dividing them by the mean of the three highest fluence points. Black squares indicate means with SE for three independent experiments with two Δphr1 lines each (separate data for each are also indicated). The lines are nonlinear least-squares fits to an exponential model, calculated as described previously (5). Statistics for the least-squares fits are given in Table 1. The black line is the fit to the combined Δphr1 data, and the gray line is a least-squares fit to phr1 fluence response curves from three independent experiments on the WT. (C) Real-time PCR data for RNA samples from the same experiments. The y axis indicates the transcript abundance of phr1 (WT) or gfp (Δphr1) relative to the abundance of gpdh transcript in the same samples and to the signal at the highest fluence, calculated as $$mathtex$$\(2^{{-}{\Delta}{\Delta}C_{T}}\)$$mathtex$$.

Quantitative reverse transcription-PCR was also performed (Fig. 6C), and the exponential fit for deleted strains clearly shows a shift to lower fluences than those for the WT; however, the shift in light sensitivity appears to be somewhat smaller than the one deduced by quantification of the hybridization signals on Northern blots. k_1/2 values for this experiment were as follows: for Δphr1 strains 3.20 and 3.3, 20.45 μmol m⁻²; and for the WT, 43.38 μmol m⁻².

We then compared the fluence response curves for phr1 induction in WT and MC strains (Fig. 7A). The k_1/2 values of MC strains were shifted to lower fluences than those for either the WT or the control strain (Fig. 7B; Table 1) carrying only the cotransformation vector. The k_1/2 value for a strain carrying a copy of the cotransformation vector, 79.6 μmol m⁻², was similar to that for the WT (82.4 μmol m⁻²). The strains with the highest copy number, MC6 and MC7, require less light for phr1 induction than the control, with k_1/2 values of 48.2 and 32.0 μmol m⁻², respectively (Table 1). Analysis of the linear range of the fluence response curves by linear regression also confirmed that this difference is significant (Fig. 7C). These strains have increased sensitivity to light for phr1 induction, but their change in light requirement was less than that for Δphr1 strains.

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FIG. 7.

Fluence response curves for the induction of phr1 in MC lines. (A) Representative set of Northern blots. (B) Fluence response curves. Circles represent the WT, and triangles and squares represent the three-copy strains MC6 and MC7, respectively. Each RNA sample was pooled from five photoinduced mycelial colonies; data are from densitometric or phosphorimager scanning of the RNA blot hybridizations from five, five, and two independent experiments for the WT, MC6, and MC7, respectively. The two scanning methods gave results that were identical within the variability. Data analysis was done as described in the legend to Fig. 6. Statistics for the least-squares fits are given in Table 1. (C) The linear range of the same data was plotted. Error bars indicate standard errors of the means for independent experiments, and the lines are linear regression plots.

These results suggested that PHR1 autoregulates its photoinduction, and we propose that this could occur through interaction with the WC light regulatory complex. To test whether this hypothesis might apply to genes other than PHR1 itself, we studied the photoinduction of the recently described blue light-regulated (blu) genes in Trichoderma atroviride (39). Photoinduction fluence response curves for blu6, blu8, blu16, and blu17 were determined with an overexpresser and a deleted strain (Fig. 8A). A decreased expression level of blu genes was observed in transgenic strains and was more remarkable in the deleted strain, at two to five times less than that in the WT strain. This effect varies between the different blu transcripts. Analysis of fluence responses curves shows that photoinduction of blu6, blu8, and blu17 in transgenic strains requires more light than that in the WT strain (Fig. 8B).

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FIG. 8.

blu gene photoinduction is altered in phr1-overexpressing and -deleted strains. (A) Photoinduction fluence response assays of phr1, blu6, blu8, blu16, and blu17 were carried out with the overexpresser strain MC7, the Δphr1 knockout strain 3.20, and the WT strain by Northern blot hybridization. The fluence of the light pulse is indicated for each lane. (B) Fluence response curves. Data are from densitometric scanning of the RNA blot hybridizations, normalized using gpdh transcript levels in the same blots.

A blue light-UVA photoreceptor is involved in phr1 induction.The UVA and blue light induction of phr1 is consistent with the photoconidiation action spectrum (18). A blue light-UVA photoreceptor is thus the sensor for phr1 induction, and there is genetic evidence that this photoreceptor is BLR1/BLR2, the WC complex in Trichoderma (7). In contrast, the wavelength dependence of phr1 induction differs markedly from the photoreactivation action spectrum. UVA is the most effective light for photoinduction, while in photorepair it accounts for 50% of the maximum. In the visible region, light effectiveness for photoinduction extends to 550 nm, while in photorepair it extends only to about 430 nm. Moreover, blue light is more effective than violet light for photoinduction (Fig. 1), while the opposite occurs in photorepair, with blue light being almost ineffective (41). In goldfish cell cultures, likewise, visible light effectiveness for photoinduction of photolyase gene expression extends beyond the photorepair range (37). If a flavoprotein is responsible for photolyase photoinduction, this difference suggests that oxidized, and perhaps semiquinone, flavin states are active.

PHR1 plays a major role in photorepair in Trichoderma.Blue light responses and DNA repair may have helped primitive organisms to evade the stress imposed by a light environment that includes damaging shorter wavelengths of UV (19, 51). Photolyase may be particularly important for Trichoderma, which lacks photoprotective carotenoids, whereas in Neurospora, blue and UVA light induce carotenoid synthesis. The complete loss of photoreactivation in Δphr1 mutants indicates that phr1 is the major component of the photorepair system in T. atroviride. The modest increase in photoreactivation capacity in PHR1-overexpressing strains suggests that the photolyase concentration is already nearly saturating for photorepair in the WT. The lack of increase in UVC survival rates for those strains suggests that either PHR1 does not contribute to dark DNA repair systems or our current method could not measure it.

Photosensory role of PHR1.Hypersensitivity to light for phr1 photoinduction in the Δphr1 and MC strains suggests that PHR1 may regulate the light input pathway for its own induction. The higher hypersensitivity to light in Δphr1 strains than in MC strains, i.e., fivefold against twofold, suggests that PHR1 may negatively regulate its own light input pathway. A gene dose effect can explain why MC strains did not show a shift to higher fluences in photoinduction. Another explanation could be that PHR1 has a more complex role than a simple negative regulator of its light input pathway.

The molecular mechanism by which PHR1 could modulate its own expression is unknown. Photolyase binds undamaged DNA with a low affinity, but the possibility of direct interaction with regulatory sequences, including those in its own promoter, must be discarded due to the lack of specificity needed for the induction of particular genes. The region upstream of phr1 (5) contains elements, such as APE and GATX boxes, similar to those found in the Neurospora gene promoters that are regulated by light through the light-responsive transcriptional activators WC1 and WC2 (12, 15, 22, 47). The Trichoderma BLR1 and BLR2 proteins (WC1 and WC2 homologs) are required for photoinduction of phr1. A BLR1 and BLR2 complex is thus likely to be the photoreceptor providing the major component of blue light and UVA regulation of phr1 (7). PHR1 might interact with the BLR complex to negatively regulate its own expression. The decreased expression levels of blu genes and their decreased sensitivity to blue light in phr1-overexpressing and phr1-deleted strains could also be explained by an interaction of PHR1 with the WC complex. Such a PHR1-WC complex interaction might be reflected in repression of the phr1 promoter in the dark and by a positive action on photoinduction of blu transcript accumulation.

Is the same PHR1 functional in photorepair and photoregulation? The difference in action spectra for photoreactivation and photolyase induction and the fact that photoreactivation requires 3 orders of magnitude more light than photoresponse led us to postulate that a specific biochemical state of PHR1 provides a photosensory function. Light effectiveness for photorepair extends to 430 nm, while in photoresponse it extends to 550 nm. If a flavin from PHR1 is involved somehow in photolyase photoinduction, this difference suggests that oxidized, and perhaps semiquinone, flavin states are more important for photoinduction than the reduced state privileged for photorepair. The MTHF chromophore seems dispensable for E. coli photolyase activity, while it appears to be more important in V. cholerae cryptochrome function. MTHF is stably attached in VcCry1, and moreover, the energy transfer from MTHF to FAD in VcCry1 is four times faster than that for E. coli photolyase (42). The MTHF chromophore of PHR1 might therefore have a more important role in photosensing than in photorepair.

Evolutionary and structural considerations for a photolyase photosensory role.The ancestral gene of the photolyase-photoreceptor family is CPD photolyase. Considering that 6-4 photolyases from animals and plants are very close to animal cryptochromes, it was proposed that the ancestors of plant and animal cryptochromes diverged before plants diverged from animals (26, 31). In this evolutionary model, plant and animal cryptochromes arose independently by repeated evolution (8). Evolution of the circadian clock would have led to a time-keeping mechanism in which light input is controlled by cryptochrome, even in large metazoans that were at less direct risk of UV damage. Finally, in mammals, cryptochrome became part of the clock oscillator, having a central but light-independent role in the negative feedback loop of the clock (20). The recent identification of prokaryotic cryptochromes belonging to the DASH group has brought up to date the initial proposition that cryptochromes diverged from photolyases before the origin of eukaryotes (6, 28, 49). Thus, of the five main subfamilies, the cluster including 6-4 photolyases and animal cryptochromes and the new DASH cryptochrome group have both photorepair and sensory functions. The plant cryptochrome cluster, in contrast, has only sensory functions, while the class I and class II CPD photolyases are known only to have DNA repair functions. The structure of cryptochromes resembles that of photolyases, and indeed, a recently determined cryptochrome structure (6) is similar to that of E. coli photolyase. Thus, it would be surprising, from the evolutionary point of view, if photolyase has no protein interaction partners. Drosophila dCRY exhibits a strong, light-dependent interaction with its partner in circadian transcriptional regulation, TIMELESS (9); the ability to interact with another member of the transcriptional regulatory complex persists (and becomes independent of light) when the C-terminal extension is deleted (40), leaving a protein which should have virtually the same structure as photolyase.

The N-terminal “tail” of fungal photolyases as a putative photosensory domain.A C-terminal domain extending 30 to 200 amino acids beyond the region of high homology with photolyases characterizes most cryptochromes and seems to be important for nucleus-cytosol trafficking and protein-protein interaction. The C-terminal “tail” is sufficient to mimic the action of light in plants (50). In Drosophila cryptochrome, it is responsible for light sensitivity, as loss of the C-terminal “tail” results in a constitutive “on” phenotype with respect to its ability to bind to its partner(s) in the circadian transcriptional regulator complex (40). Fungal photolyases and DASH cryptochromes lack this specific additional sequence, but they do extend 70 to 140 amino acids towards the N terminus. In fungal photolyases, this N-terminal extension contains the putative mitochondrial and nuclear localization signals, while in Cry3, a DASH cryptochrome, it contains a transit peptide for localization in chloroplasts and mitochondria. This N-terminal extension might confer some photosensory properties (Fig. 9A). Alignment of the N-terminal 140 amino acids of visible light-regulated fungal photolyases, namely, those of T. atroviride (5), N. crassa (Berrocal-Tito, unpublished results), and Fusarium oxyosporum (2), shows that this region is very well conserved (Fig. 9B). A standard BLAST analysis of this N-terminal region of T. atroviride photolyase does not show any specific homology, while an RPSI-BLAST analysis against the BLOCKS database (25) shows significant alignments with presenilin, amphiphysin, and Sac3/GANP proteins, which have in common a protein binding ability, suggesting an interacting role for this region. In addition, putative phosphorylation sites for either cyclic AMP- or cyclic GMP-dependent protein kinase, protein kinase C, and casein kinase II are found by comparison to the PROSITE database, at amino acid positions 44 to 48 of PHR1. In class I photolyases, light excites MTHF, and this energy excitation is transferred to FAD, perhaps promoting a conformational change in PHR1 that could result in N-terminal modification, for example, phosphorylation.

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FIG. 9.

Comparison of N-terminal regions of light-regulated fungal photolyases. (A) Schematic comparison of fungal photolyases and cryptochromes. (B) Alignment of the 140 N-terminal amino acids of light-regulated fungal photolyases. The underlined region of PHR1 shows homology with presenilin and amphiphysin, and the heavy line indicates a phosphorylation site. E. c., E. coli; At, A. thaliana; Nc, N. crassa; Fo, F. oxyosporum; Ta, T. atroviride.

To explain the findings reported here, we propose the following model (Fig. 10), which can be tested by further molecular genetic and protein interaction experiments. In this model, PHR1 interacts with or is part of a transcriptional regulatory complex. To be part of this complex, PHR1 must have specific photochemical properties, resulting, for example, from a posttranslational modification of the N terminus. PHR1 present in the dark would exert an inhibitory effect on photoinduction by recruiting an inhibitory protein to the complex; light relieves this repression. This would be completely removed in null mutants, leading to the observed increase in light sensitivity. A weak gfp hybridization signal was detected even in the dark in some, but not all, Northern blots (just visible in Fig. 6A) and was more significant in real-time PCR experiments (Fig. 6C). This suggests that a basal level of PHR1 might enhance binding of the repressor already present in the dark. Overexpression might lead to accumulation of excess free PHR1 in the nucleus, partially titrating away the inhibitory protein. Thus, the light-dependent interaction of cryptochrome/photolyase family members with transcriptional regulators may predate the divergence of cryptochromes from photolyases.

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FIG. 10.

Photolyase light-dependent autoregulation model. Circles, blue light regulators (BRL) 1 and 2; gray box and solid line, phr1 promoter and coding regions; black and gray polygons, DNA photolyase (PHR1) with different photochemical properties; rectangle, putative repressor; lightning, exogenous light. Photolyase would act as a modulator of its own transcription through interaction with the BRL complex, recruiting a repressor to the complex in the dark. Light relieves repression by an unknown mechanism, resulting in activation of phr1 transcription, which is dependent on BLR1 and BLR2. In addition, light activates PHR1 for DNA repair.