1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Phytochrome A (phyA), the most versatile plant phytochrome, exists in the two isoforms, phyA′ and phyA′′, differing by the character of its posttranslational modification, possibly, by phosphorylation at the N-terminal extension [Sineshchekov, V. (2010) J. Botany 2010, Article ID 358372]. This heterogeneity may explain the diverse modes of phyA action. We investigated possible roles of protein phosphatases activity and pH in regulation of the phyA pools' content in etiolated seedlings of maize and their extracts using fluorescence spectroscopy and photochemistry of the pigment. The phyA′/phyA′′ ratio varied depending on the state of development of seedlings and the plant tissue/organ used. This ratio qualitatively correlated with the pH in maize root tips. In extracts, it reached a maximum at pH ≈ 7.5 characteristic for the cell cytoplasm. Inhibition of phosphatases of the PP1 and PP2A types with okadaic and cantharidic acids brought about phyA′ decline and/or concomitant increase of phyA′′ in coleoptiles and mesocotyls, but had no effect in roots, revealing a tissue/organ specificity. Thus, pH and phosphorylation status regulate the phyA′/phyA′′ equilibrium and content in the etiolated (maize) cells and this regulation is connected with alteration of the processes of phyA′ destruction and/or its transformation into the more stable phyA′′.


far-red light


extent of the phototransformation of the initial form of phytochrome into the first photoproduct


high-irradiance response

λa, λe, λmax

wavelengths of the actinic and excitation light and of the maximum of phytochrome fluorescence


low fluence response


the first photoproduct of the phototransformation of phytochrome in its red form stable at 77–85 K

OA and CA

okadaic and cantharidic acid




total phy content


red-light absorbing form of phy


far-red-light absorbing form of phy


red light

PP1 and PP2A

protein phosphatases 1 and 2A


phytochrome kinase substrates 1–4

phyA and phyB

phytochromes A and B

phyA′ and phyA′′

subpopulations of phyA


very low fluence response


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

One of the major steps forward in phytochrome (phy) research was the discovery of its structural and functional heterogeneity (see Ref. [1] for a review). The phytochrome family consists of a small number of phy members (phyA–phyE in Arabidopsis), the major ones being phyA and phyB. The light stable phyB shows classical characteristics of phy light sensing at low light fluences (the red-induced/far-red-reverted low fluence responses, LFR). phyA, which is the predominant phytochrome in etiolated seedlings, whose major fraction is light labile, shows a quite different behavior: it promotes irreversible effects in the whole range of its absorption spectrum under very weak light (very low fluence responses [VLFR]) and under high fluence rates with the maximum activity in the far-red region (high-irradiance responses, HIR). Along with these well established properties of phyA, it may perform as well the LFR functions characteristic for phyB [2-5]. In particular, active phyA is translocated to the nucleus with the help of FHY1 and FHL, possibly via their red (R)/far-red (FR) light reversible phosphorylation [6-8]. In the context of the different types of the phyA photoresponses, the differentiation between cytoplasmic and nuclear phyA functions is relevant (as reviewed in Ref. [9]). Not all the phyA translocates to the nucleus, part of it remains active in the cytoplasm. Mediation of root phototropism by phyA together with phyB under R [10, 11] and modification of gravi- and phototropism [12-15] can be attributed to the cytoplasmic fraction [16].

These different functions and properties of phyA were explained by its heterogeneity (see Refs. [2, 17-19]). With the use of low-temperature fluorescence spectroscopy two phyA types, phyA′ and phyA′′, were detected in wild-type mono- and dicotous plants and their phy mutants. They differ by spectroscopic and photochemical properties, and their content depends on plant species and tissues and on physiological conditions. One of them, phyA′, predominates in growing tissues and is light labile. It has the emission (absorption) maximum at λmax = 687(673) nm at low temperatures, and is efficient in Pr [RIGHTWARDS ARROW] lumi-R photoconversion at 77–85 K (Pr′ photochemical type). In contrast, phyA′′ is a minor species more stable in the light and its concentration does not change significantly with tissue type. It has λmax = 682–683 (668) nm and is ineffective in the Pr [RIGHTWARDS ARROW] lumi-R conversion at low temperatures (Pr′′ photochemical type). phyA′ is responsible for de-etiolation under FR (HIR and VLFR) whereas the relatively light stable phyA′′ is likely to be active under R and could be functional together with phyB throughout the plant's life cycle. phyA′′ suppresses the action of phyA′ and the regulation of the ratio between phyA′ and phyA′′ appears to be important for phyA function. The notion of the two different functional phyA populations is gaining support in the recent photophysiological observations [5]: phyB phyC double mutant (of rice) containing only phyA displayed clear R/FR reversibility in pulse irradiation experiments, indicating that there is a fraction of phyA which together with phyB can mediate the low-fluence response for gene expression. phyA′ and phyA′′ are the products of one and the same gene and differ only by posttranslational modification at the N-terminus [20]. This modification is likely phosphorylation because the equilibrium between phyA′ and phyA′′ is connected with phosphorylation/dephosphorylation of the pigment and with its possible interaction with probable transphosphorylation substrates, such as PKS1 and PKS2 and CRY1 [18, 21].

phyA was shown to be itself a phosphoprotein and a light-regulated kinase [22, 23]. Under illumination, it autophosphorylates and transphosphorylates some partners [24, 25]. Cryptochromes, CRY1 and CRY2, are also phosphorylated by phyA in vitro and the effect for CRY1 is light dependent in vivo [26]. AUX/IAA proteins can be phosphorylated by phyA too [27]. In the dark, phyA is phosphorylated at Ser8 (in vivo) and Ser18 (in vitro) residues and, upon illumination, phosphorylation at the C-terminal part (at Ser598) takes place (in vivo) [28]. From experiments on mutated phyA, it is known that phosphorylation of phyA modulates its activity [25, 29-31]. These experiments have shown that phytochrome phosphorylation at Ser598 is involved in inhibition of its action by interfering with the pigment's interaction with its putative signal transducers (NDPK2 and PIF3). In line with this, it was shown that a Type-5 protein phosphatase (PAPP5) specifically dephosphorylates phytochrome in its biologically active Pfr form and enhances phytochrome-mediated photoresponses [32].

Recently, Han et al. [33] identified Ser8 and Ser18 in the 65 amino acid N-terminal extension (NTE) region of oat phyA as being the autophosphorylation sites. Autophosphorylation site phyA mutants (in phyA deficient Arabidopsis thaliana) formed sequestered areas of phytochrome (SAPs) in the cytosol much more slowly than did wild-type phyA and the mutant pigment degraded at a significantly slower rate than wild-type phyA under light conditions. At the same time, these mutants had an increased activity. These results suggest that the autophosphorylation of phyA plays an important role in the regulation of plant phytochrome signaling through the control of phyA protein stability. In the context of the present discussion of the two phyA types, which may differ by phosphorylation at the N-terminus and by the character of their functional activity, it is of particular interest that phyA phosphorylation at the N-terminus extension may serve as a means of discrimination between different modes of the phyA action [3].

The possible implication of the phosphorylation/dephosphorylation status of the cell in the formation of the two phyA species prompted us to turn to the investigation of effects of agents changing phosphorylation of phyA on the phyA′/phyA′′ equilibrium. These are specific phosphatase inhibitors modifying phosphatase/kinase activity in the cell and affecting phyA indirectly in vivo as well as animal and bacterial phosphatases which could directly alter phosphorylation state of phyA in in vitro systems—in crude extracts and pure phyA solutions. Another major factor which may affect either phytochrome molecular properties directly [34, 35] or indirectly via modification of biochemical processes in general [36] is pH of the phyA environment.

Okadaic (OA) and cantharidic (CA) acids—structurally unrelated compounds—were shown to be potent and specific inhibitors of protein phosphatases 1 and 2A (the PP1 and PP2A types, respectively), which act at the Ser/Thr residues (see Refs. [37, 38]) and the literature cited therein). These phosphatases are involved in a number of regulation processes including activation of metabolic enzymes, ion channel regulation, gene expression and developmental processes. Phosphorylation is partly responsible for maintaining high rate of microtubule dynamics and okadaic acid was shown to increase it [39]. In particular, root and hypocotyl formation, gravity response and auxin transport are modified in Arabidopsis mutants with defects in the gene encoding for one of the subunits of PP2A [40, 41]. In the context of phytochrome research, it is of importance to note that okadaic acid influences light-induced effects. It is shown to block chlorophyll accumulation in etiolated maize leaves and expression of a number of photosynthetic genes. From these and other experiments, it was suggested that PP1/PP2A is required for transmitting light signals and light-dependent gene activation in plants [37].

Proceeding from the above considerations, we have carried out experiments in two major directions—we tried to follow effects of changes in (1) phosphatase/kinase activity and (2) pH variations on the phyA′/phyA′′ equilibrium in three systems of different complexity—intact etiolated maize tissues, crude extracts from them and pure phyA solutions. Destruction of phyA′ and/or its conversion into phyA′′ were observed upon the action of specific PP inhibitors (against PP1 and PP2A) in shoots but they were lacking in roots. On the other hand, strong effects of pH variations on the phyA′/phyA′′ equilibrium were found in vivo and in extracts. These data suggest that the phosphatase activity of the cell and its pH may modulate destruction of the labile phyA′ form and/or its conversion into the more stable phyA′′ already in the dark.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Conditions of plants growth and preparation of tissue samples and extracts

Zea mays L. plants of the following varieties were used: Kubanskaya and LG 32.26 Lukas, Limagrain Nickerson GmbH (labeled as LG 32.26 in this work). The seedlings were grown at 20–23°C on filter paper soaked with tap water in total darkness. The samples measured were tips of coleoptiles or roots and mesocotyl segments. The length of the samples and the age of the seedlings varied depending on the character of the specific experiment (see below). In a number of experiments, etiolated seedlings of Arabidopsis (without cotyledons) were also used. All the manipulations with the samples were carried out under photochemically safe green light (fluorescent lamps with bandpass green filters or green light diodes LG 3330/HV12).

In the experiments on phyA state in developing roots, seeds of maize imbibing for 24, 48 or 72 h were taken and samples for measurements were root tips (1 ± 0.3 mm) at three different stages of their development [42]: stage 1—radicle protrusion and initial growth, when the total root length (L) is 0.7–2 mm, stage 2—cell elongation in the root tip (L = 7–10 mm) and stage 3—cell differentiation in the root tip (L = 15–30 mm). These changes in the root development correlate with a certain extent with variations in the cellular pH in root tips (see below).

Material for investigations in the experiments on the effect of PP inhibitors was tips of coleoptiles (L = 27 ± 5 mm) and roots (L = 13 ± 1 mm) of 3 to 4-day-old and 2 to 3-day-old etiolated seedlings of maize, respectively, and also crude extracts from them. Extracts from Arabidopsis seedlings (4-day-old) were also used. Treatment of plant tissues with inhibitors was carried out using specific phosphatase PP1 and PP2A inhibitors okadaic and cantharidic acids—both at concentrations 0.5 μm. Treatments with the inhibitors were carried out in the dark according to the following schemes—10 min, 4 h and 4 + 12 h in tap water to follow delayed effects. It should be noted that the scheme “4 h inhibitor + 12 h H2O” was used earlier by Sheen [37]: 4 h treatment of maize shoots in 0.5 μm solution of OA in the dark was followed by 12 h light exposition in water. The author found out that such a procedure drastically hampered the greening process believed to be a result of inhibition of the expression of light-inducible genes. At the same time, there was no inhibition of general cellular metabolism even at a higher OA concentration (1 μm). This implies that the treatments with the inhibitors were not destructive for the samples. In this work, tissues' vitality was controlled with fluorescein and/or eosin dyes. To exclude a possible harmful effect of the inhibitors, dead tissues (roots treated for 4 h with CA 100-fold more concentrated than that used in the experiments or with 70% ethyl alcohol for 20 min) were used as a control. The so treated roots looked quite different from the intact ones and possessed strikingly different spectroscopic characteristics, belonging probably to the degraded phyA and other molecules. This demonstrated that the experimental conditions chosen allowed observation of viable tissue.

The seedlings were treated either as a whole intact plant with a root in solution or as a shoot cut at the level of coleoptile or coleoptile + mesocotyl with the lower end put into solution and upper part above the solution (in the case of coleoptile and mesocotyl samples). The root and coleoptile tips and mesocotyl parts were cut off for sample preparation after incubation with inhibitors at 28°C in darkness.

To follow pH change during root development from radicle protrusion to the root formation, we used root tips of 1–1.5 mm (roots: 48 h old, total length from 1 to 30 mm). They were incubated in fluorescein diacetate (FDA) solution (10 μm) for 15 min at 26°C and then rinsed for 1 min in distilled water. Fluorescence spectra of the treated roots (six root tips in a sample) were measured upon excitation at λ = 488 nm and λ = 450 nm. Root tips incubated for 15 min in water were used as a control. Two independent samples were used for each data point. The pH in the root tips was calculated from the FDA fluorescence and its calibration curve.

To prepare a sample, two maize coleoptile tips (3–5 mm long), three root tips (1–2 mm long) and from 10 to 25 Arabidopsis hypocotyls were taken and glued with water/glycerol (50/50 vol/vol) mixture, which is used as a cryoprotector, to a Plexiglas plate. Excess water/glycerol mixture was taken away with a piece of filter paper. The plate with the sample was either fixed in a cryostat (at the bottom of a hollow copper cylinder filled with liquid nitrogen and put into a transparent Dewar flask; at 85 K) or immersed directly into liquid nitrogen (77 K). The procedure of sample preparation for the measurements was essentially the same as in [43].

In a number of experiments, crude extracts from maize and Arabidopsis seedlings and pure phyA solutions were also used. In the experiments on OA and CA effects on phyA in extracts from maize roots or coleoptiles, three tips were usually taken and homogenates were prepared in a cooled down to 0°C mortar in 40 μL of 50 mm Tris-HCl buffer (pH 8) which contained 5 mm EDTA. The extract was transferred on a plate of the sample holder and frozen down to 77–85 K. (It should be noted that under these conditions the proportion of the two phyA types in the extracts did not differ significantly from that in native tissues suggesting that the state of phyA in them was not modified.) For a control, tips of roots kept in distilled water and tips of coleoptiles and parts of mesocotyl without treatment were taken. For experiments with lambda protein phosphatase and alkaline phosphatase action on phyA, 1 g of etiolated tissue was homogenized in the HEPES buffer (pH = 7.6) or Tris buffer (pH = 7.9) in the proportion 1:1 at 0°C. The extract was centrifuged at 4°C for 5 min at 13 200 g (Eppendorf 5415R). The supernatant was incubated in a 1.0 mL Eppendorf tube for 1 h at 30°C (for λ-PP) or 35°C (for CIAP).

Measurements of extracts and pure phytochrome solutions were carried out in a small (250 μL) PCR tube or as a droplet on the surface of the Plexiglas plate fixed similarly either in the cryostat or directly in the Dewar flask. For cryoprotection and better light-scattering conditions, glycerol was added to the extracts and solutions to obtain a 1:1 vol/vol ratio mixture.

In the experiments on pH modulation, a linear correlation between pH in Tris buffer and pH in crude extracts was initially obtained (Fig. 1). From this dependence such a pH of Tris buffer was chosen so that the necessary pH value in crude extract in the pH range from 6.5 to 8.5 was obtained.


Figure 1. Correlation between pH in Tris buffer and pH in crude extracts (CE) from etiolated maize coleoptiles prepared with the respective buffer.

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Arabidopsis phyA expression in Pichia pastoris and E. coli and purification

phyA full-length coding DNA sequence (CDS) as well as the CDS coding for a 619 amino acid sensor domain part were amplified by PCR with oligonucleotides MZ349 (CTAGGGATCCACCATGGGCTCAGGCTCTAGGCCGACTCAGTCC), MZ371 (GAGGGCCCATGGTGATGGTGATGGTGGCCCTGAAAATAAAGATTCTCCTTGTTTGCTGCAGCGAGTTCCGCAGTG) and MZ369 (GAGGGCCCATGGTGATGGTGATGGTGGCCCTGAAAATAAAGATTCTCTTCTAGTTCTTGTATACCATCAAT), respectively. PCR fragments were NcoI and NotI digested and ligated into NcoI and NotI digested pPICZ (Invitrogen). Positive clones, pPICZphyA and pPICZphyAΔ3, were transformed by electroporation into Pichia strain X-33 (Invitrogen) and yeast clones with multiple insertions were selected with 1500 μg mL−1 Zeocin following manufacturer's instructions. Resistant clones were screened for expression level and clones with high expression level were used for shaken flask culture. Cultures were grown in MGYH (Invitrogen) to OD 1–2 where the cells were harvested by centrifugation. Expression was induced by resuspension of the cell pellet in MMH media (Invitrogen) and was carried out for 3 days with MeOH supplementation every 12 h up to a concentration of 0.8% (vol/vol). Cells were harvested by centrifugation and resuspended in 3 vol. extraction buffer (50 mm Tris HCl pH 7.8, 10 mm EDTA, 150 mm NaCl). Cells were broken in liquid nitrogen with an Ultraturrax (ArtLabortechnik, Germany) at full speed for 20 min. The resulting snow was thawed, mixed with excess PCB [44] and immediately centrifuged for 60 min at 4°C at 40 krcf. The supernatant was ammoniumsulfate-precipitated with a final NH4SO4 concentration of 40% (wt/vol) for 12 h. The precipitate was harvested by centrifugation for 10 min at 4°C and 20 krcf and resuspended in AB-buffer (10 mm Imidazole, 50 mm Tris HCl pH 7.8, 150 mm NaCl) and subjected to Ni-NTA affinity purification (Qiagen). Bound protein was eluted with EL-buffer (150 mm Imidazole, 50 mm Tris HCl pH 7.8, 150 mm NaCl) and rebuffered by NAP-desalting columns (GE) into extraction buffer. The final solution was concentrated in an Amicon stirred cell (molecular weight cut off 20 kDa) with pressurized nitrogen.

For expression in E. coli, the above employed plasmids were digested with NcoI and NotI which liberated the phyA full length and sensor domain coding CDSs. Those were ligated into NcoI and NotI digested pET28c (Novagen). The resulting plasmids were cotransformed with plasmid p171 [45, 46] into E. coli Rosetta cells (Novagen). Positive clones were selected and grown in LB to an OD of 0.5. Expression was induced with 1 mm Isopropyl β-D-1-thiogalactopyranoside for 8 h at 18°C. The small fraction of soluble protein was purified as above except a French press was used to break the cells at 4°C with three passes. The integrity of the proteins thus obtained was shown by Western blot detection of full-length phyA and Coomassie stain detection of purified full length and sensor domain part of phyA.

Fluorescence and conventional light microscopy

To follow possible structural effects of the phosphatase inhibitors, viability of the cells after the treatment was determined with the use of fluorescence microscopy (fluorescence microscope Zeiss Axiovert 200) after the FDA staining (20 min in 0.005% wt/vol solution) as described (see Ref. [47] and the literature cited therein). The samples were prepared according to [48, 49]. For light microscopy, semifine sections of root tips were obtained essentially as described in [50] with some modifications, stained with toluidine blue and photographed with CAMEDIA digital C4040 camera. We did not observe any visual difference of the cells taken from the control or experimental samples during OA treatment (≤12 h) which could be associated with cell damage.

Fluorescence spectroscopy of phytochrome. Acquisition of phyA characteristics from the raw emission spectra of the samples

The experimental approach was essentially the same as that employed earlier in our research [17-19, 43, 51]. Briefly, we used low-temperature (77–85 K) fluorescence of phy for its in situ assay. Fluorescence emission spectra of phy were taken in its Pr state (F0), after transformation into lumi-R upon saturating red illumination at 85 K (F1—state of photoequilibrium between Pr and lumi-R at 85 K) and after partial phototransformation into Pfr at 273 K (F2—state of photoequilibrium between Pr and Pfr) and again after phototransformation of Pfr into Pr upon illumination with FR light at 273 K (F3—state of photoequilibrium between Pr and Pfr under FR at 273 K). Full reversibility under these conditions was taken as a firm proof that the emission belonged to phyA. Fluorescence intensity of phy in its initial Pr form (F0) was taken as a measure of the total phy content in the sample ([Ptot] in rel. units) whereas its relative changes under saturating red illumination at low temperature, as the extent of the Pr [RIGHTWARDS ARROW] lumi-R conversion (γ1). The latter value is a measure of the samples relative content of the phyA′ species, which is active in the low-temperature Pr [RIGHTWARDS ARROW] lumi-R photoconversion (its individual γ1′ = 0.5) and of the phyA′′ species, which is inactive in this photoconversion (γ1′′ = 0). The evaluation of the total phyA content and the calculation of the ratio of its two native pools (phyA′ of the Pr′ type and phyA′′ of the Pr′′ type) were done according to the procedure described in [43]. In these evaluations, phyB, which also belongs to the Pr′′ photochemical type, was not taken into account because of its relatively low content (less than 5%).

The phototransformation of Pr into lumi-R was carried out upon R illumination at 85 K, whereas all fluorescence spectra were taken at 77 or 85 K, depending on the specific experimental setup. The rise of the temperature from 77 to 85 K is needed to increase the rate of the photoconversion of Pr into lumi-R and bring the temperature of the photoconversion close to that for which the individual γ1 values of phyA′ and phyA′′ were determined earlier [43, 51]. It should be noted that the extent of the Pr [RIGHTWARDS ARROW] lumi-R conversion did not change considerably with the temperature in the range 85 ± 5 K [52] so the precision of the determination of the sample temperature in these measurements did not affect the obtained γ1 values. The photoconversion Pr [RIGHTWARDS ARROW] Pfr can be achieved only at ambient temperatures.

Experimentally, two raw emission spectra were usually taken to characterize phy in the sample: one immediately after freezing at 77–85 K (Fig. 2a, curve 1) and another of the same sample after saturating illumination and conversion of the Pr form into the initial photoproduct stable at low temperatures, lumi-R (Fig. 2a, curve 2). The two raw spectra of the sample under investigation were corrected for the background fluorescence by the subtraction of the spectrum of the reference sample (mature roots at their base after saturating red light illumination to convert Pr into Pfr in case of maize (Fig. 2a, curve 3) or hypocotyls of phyA-less mutant in case of Arabidopsis). The sample of mature FR-illuminated maize roots did not practically contain phy in its Pr form and could be used as a spectrum of the background light (reference spectrum). Earlier experiments with phyA phyB double mutants of Arabidopsis and pea [53, 54] proved the validity of such an approach by showing that the background spectrum of the wild type thus obtained and the spectrum of the double mutant virtually coincided.


Figure 2. Low-temperature (77 K) fluorescence emission spectra (exciting light from He–Ne laser, λex = 632.8 nm, fluence rate reduced to 0.5 W m2) of (a) etiolated maize (LG 32.26 var.) mesocotyls at their base (raw spectra) and of phytochrome in this tissue (difference spectra) (b). The spectra were taken after freezing the sample in the dark at 77 K [1] and after its warming up at 85 K and saturating illumination with full laser light at this temperature (λa = 632.8 nm, I = 6.5 W m2, 10 min) partially transforming the initial red-absorbing phytochrome form, Pr, into the first stable at low temperatures photoproduct, lumi-R [2]. Curve 3 is a fluorescence spectrum of maize tissue which contains virtually no phytochrome in its Pr form—old maize roots at their base taken after saturating red illumination (I = 6.5 W m2, 5 min) at room temperature to convert phytochrome in its Pr form into Pfr. It is taken as a spectrum of background emission of the sample. The spectra of phy in (b) were obtained by subtracting spectrum 3 from spectra 1 and 2 (the spectra in this work were not corrected for the spectral sensitivity of the spectrofluorimeter.).

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After normalization of the raw spectra and the reference spectrum to unity at 657–660 nm, where the input of phy is less than 3–5%, the reference spectrum was subtracted from the raw spectra to give the spectra of phy in the sample (Fig. 2b). From them, we obtained position of the emission spectrum (λmax), [Ptot] in rel. units and γ1, which is calculated as the difference in the intensity of the two spectra in the maximum (ΔF = F0F1) related to F0, γ1 = ΔF/F0). Knowing [Ptot] and γ1, it was easy to evaluate phyA′ and phyA′′ (+ phyB) in the sample in % and rel. units (see 'phyA native pools in etiolated maize roots and coleoptiles'). To get the phyA′ and phyA′′ parameters, five to seven independent samples were measured and the data are presented as means ± SE. The Student's t-tests (t-criterium) was used (the level of confidence of 0.05) for independent distributions. Each distribution was checked for normality using Fisher's criterion (with the level of significance value of 0.05) for each compared populations.


Three different spectrofluorimeters were used in these experiments. First, a laboratory-designed instrument based on two double-grating monochromators of the DFS types (diffraction spectrometers, LOMO, Leningrad, USSR) conventionally used for the Raman spectroscopy was employed [43]. Second, a spectrofluorimeter (Shimadzu RF-5000; Shimadzu, Tokyo, Japan) equipped with a laboratory-designed application for sample cooling as in our earlier measurements [55, 56]. It consisted of a bifurcated light guide, of which one branch was used for conducting exciting light to the sample and the second branch for directing the emission light from the sample to the entrance aperture of the emission monochromator of the spectrofluorimeter. To achieve this, the end of the emission branch was fixed to the cuvette holder at the position normally taken by the standard cuvette. The fused branch of the light guide was attached “head on” to a quartz glass rod (covered with metal), whose other end was attached to a cylindrical sample cuvette. The only difference from the former setup in the current experiments was the use of a He–Ne laser (2 mW, type PL 610 Klasse 3b; Polytec, Waldbronn, Germany) instead of the Shimadzu RF-5000 exciting monochromator. Also, instead of the cuvette, the sample was attached directly to the glass rod to allow for a better light collection. The laser was used as a source of the exciting/actinic light at λ = 632.8 nm with its beam focused on the entrance aperture of the excitation branch of the light guide. In the case of exciting mode, the laser was supplied with three interference filters (transmitting at 633 nm), which were put in to cutoff pumping light interfering with spectra measurements and to reduce the energy fluence rate of the light from 6.5 W m−2 (at the plane of the sample) to 0.5 W m−2 (more than 10-fold) so that it did not produce noticeable photoconversion of the Pr form of phytochrome during spectra measurement. For the actinic illumination of the sample with full laser light, the filters were taken off. To cut off exciting light, a red plastic filter with λ ≥ 635 nm (lucite 3 mm, type 502; Röhm GmbH, Darmstadt, Germany) was placed at the entrance of the analyzing (emission) monochromator. And finally, a third spectrofluorimeter (FluoroMax-4; HORIBA-Scientific, France) with a liquid nitrogen Dewar assembly was used. In this case, the actinic light was that from the exciting monochromator at λa = 620 or 660 nm. Its intensity was reduced by neutral (18% transmission) and interference (620 nm) filters so that it did not practically produce any photochemical changes in the sample during spectra measurements at low temperatures. Red cutoff filter (transmitting at λ > 650 nm) was put at the entrance slit of the analyzing monochromator. Light-emitting diodes for red (λ = 655 nm; ELD-655) and for far-red light (λ = 740 nm; ELD-740), both from Roithner, Austria, were employed for photochemical transformation of the Pr and Pfr phytochrome forms. The far-red LEDs were equipped with a cut off filter (λ > 695 nm “black plexiglas”; Röhm).


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

In our earlier works, we observed wide variations of the phy parameters in plants depending on plant species and tissues used, developmental and physiological state of the plant, experimental conditions (illumination, for instance) etc. This reflected the presence of the two native phyA types in the plant with different spectroscopic and photochemical properties (see 'Introduction'). To understand the mechanism(s) regulating their content in the cell, we concentrated here on the complex phenomenology of the phyA pools' changes in dark-grown maize seedlings and tried to correlate them with the action of the two major regulation factors in the cell—phosphatase/kinase activity and pH.

phyA native pools in etiolated maize roots and coleoptiles

phyA and its native populations in etiolated maize seedlings were investigated by analyzing its fluorescence and photochemical parameters as a function of (1) its localization in different tissues or organs and (2) of the developmental state of the given organ/tissue. Exploring the state of phyA in growing root tips, we used traditional characterization of their development, which include radicle penetration (stage 1), cell elongation in the root tip (stage 2) and cell differentiation in the root tip (stage 3) [42]. It was found that the spectroscopic and photochemical characteristics of phyA in the root tip (ca 1 mm long) widely varied depending on the three different stages of its development. At stage 1 (total root length L = 0.7–2 mm), a rapid redshift in the position of the emission spectrum, λmax, was followed: initially, for the emerging root, λmax = 682 nm and then it reached 683.5 nm along with the root growth (Fig. 3) and further on, at stages 2 and 3, there were no changes in this parameter.


Figure 3. Low-temperature emission spectra (λex = 633 nm, normalized at the maximum) of phytochrome A in the maize root tip (Kubanskaya var.) at the initial stage 1 of its development: [1] emerging root (L = 0.7 mm), [2] root at the end of stage 1 (L = 2 mm).

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The total phytochrome content (Ptot) and the extent of the Pr [RIGHTWARDS ARROW] lumi-R conversion, γ1, were the highest at stage 1, raising, respectively, from 1.50 relative units and 0.20, when the root's length was minimal (≤1 mm), to 5.16 rel. units and 0.43, when it reached length of ca 2 mm (Fig. 4). During the following two stages the phy content steadily declined whereas the γ1 parameter revealed a more complex behavior: lower but stable values of the two parameters were registered during the second stage (L = 7–10 mm): [Ptot] = 3.39 rel. units and γ1 = 0.23, and at stage 3 (L = 15–30 mm) [Ptot] was reduced to 2.13 rel. units, however, γ1 increased to 0.33 (Fig 4).


Figure 4. Changes in the total phyA content (Ptot) (a) in the maize (Kubanskaya var.) root tip (L = 0.7 mm) and its photochemical characteristic γ1 (b) during the three stages of the root development (root length L = 1–2, 7–10 and 15–30 mm, respectively).

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These variations are interpreted, in agreement with our earlier findings, as a manifestation of the existence of the two phyA species—phyA′ and phyA′′ (see 'Introduction'). Evaluations of their content (as indicated in 'Materials and Methods') revealed that it strongly changed in a complex manner in the roots as demonstrated in Fig. 5. At stage 1, the content of phyA′ and phyA′′ are approximately equal at the beginning but along with the root elongation and total phytochrome accumulation, phyA′ concentration steadily rises and it begins to strongly dominate over phyA′′, whose absolute content, respectively, declines (Fig. 5a). Later on (stage 2), the contents of phyA′ and phyA′′ became approximately equal and then (stage 3) phyA′ again dominated over phyA′′ (Fig. 5b).


Figure 5. Content of the two native phyA populations, phyA′ (light gray) and phyA′′ (dark gray), in the tip (L = 0.7 mm) of the growing etiolated maize root (Kubanskaya var.): (a) initial stage 1 (L ≤ 2 mm) and (b) stages 1, 2, and 3 (L = 1–2, 7–10 and 15–30 mm, respectively).

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The observed pattern of the phyA′/phyA′′ distribution in developing roots confirms our previous observations [43] about the variable relative and absolute content in different tissues/organs. For example, phyA in dark-grown maize seedlings with well-developed roots and shoots revealed the following parameters: in coleoptile tips, λmax = 684 nm, F0 = 3.53 ± 0.2 rel. units and γ1 = 0.32 ± 0.02; in root tips, λmax = 683.5 nm, F0 = 2.57 ± 0.21, γ1 = 0.31 ± 0.03. According to these Ptot and γ1 parameters, the phyA′/phyA′′ proportion in these tissues were close to each other (64/36% and 62/38%, respectively), although their content was lower in the root tips (2.26/1.27 rel. units and 1.59/0.98 rel. units, respectively). In general, there was a dependence between the concentration of its native species and the total phyA content in etiolated maize tissues such that at low Ptot their content was comparable, whereas at higher Ptot phyA′ rises almost linearly with Ptot, but phyA′′ reveals early saturation (Fig. 6).


Figure 6. Dependence of the content of the two native phyA types, phyA′ and phyA′′ (open triangles and open circles and their polynomial fits 1 and 2, respectively) on the total concentration of phyA, [Ptot], in etiolated maize roots (a) and coleoptiles (b) at different stages of their development (summary data of different samples of the Kubanskaya var).

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Thus, the above phenomenology of the two phyA species in dark-grown maize seedlings indicates that they are present already in etiolated tissues, phyA′ being a variable species whereas phyA′′ is more stable and the total amount of phyA′′ reaches a saturation. It appears that phyA′′ dominates and its amount is comparable with phyA′ in roots on the very early stages of their development (Fig. 5a), but saturation of phyA′′ upon the Ptot rise is followed when Ptot reaches 1/3–1/2 its maximal values (Fig. 6a). The phyA′ concentration, on the contrary, rises almost linearly with the total phyA increase. Interestingly, a similar pattern of the phyA′ and phyA′′ is observed in coleoptiles (Fig. 6b). This and also the fact that phyA is synthesized in heterologous systems only as phyA′′ (see [20] and below) imply that phyA′ is formed from phyA′′ upon its modification, possibly, by way of phosphorylation, and there is a factor preventing its full conversion and limiting phyA′′ to a certain (ca 1/7 of maximal Ptot) relatively constant level. Assuming that phyA′′ could be a membrane-/protein-associated fraction of phyA whereas phyA′ is soluble this limiting factor could be a constant number of phyA′′-binding partners or sites. These phyA′′ molecules cannot be modified (phosphorylated?), whereas all others are converted into phyA′. Close similarity between the observed patterns of the phyA′ and phyA′′ changes with total phyA in maize roots and coleoptiles indicates that the factors and processes regulating the pools′ abundance could be of a universal character. With this in mind, we posed the question if changes in the phosphatase/kinase activity and pH could contribute to the complex regulation of phyA′ and phyA′′ in dark-grown plant tissues.

Effects of specific inhibitors of the protein phosphatases of the PP1 and PP2A types on phyA in etiolated seedlings

Short-term (up to 4 h) treatment of etiolated maize tissues (coleoptiles, root tips and mesocotyls) with okadaic acid (0.5 μm) did not produce noticeable effects on all the investigated phy parameters (Table 1). Longer exposures affected primarily the photochemical phyA parameter γ1 in coleoptiles and mesocotyls, the effect being dependent on the time of the exposures: it was relatively low after “4 h in OA + 12 h in water” and much more pronounced after “12 h in OA”. Spectroscopic characteristics were virtually identical to those of the control samples (spectra not shown), the total phy content, Ptot, remained unchanged or underwent a decline (Fig. 7a), however, the γ1 parameter dropped down by ca 2–2.5-fold (Fig. 7b). Figure 8 illustrates changes induced in phyA and its pools by OA treatment of coleoptiles. In root tips, OA does not produce any effect even after prolonged incubation: Ptot and γ1 are comparable with the intact tissues.


Figure 7. Total content of phytochrome (a) and extent of the photoconversion of its Pr form into photoproduct lumi-R, γ1 (b) in coleoptile tips and mesocotyl base of etiolated maize shoots (LG 32.26 var.)—intact tissues and okadaic acid (0.5 μm) treated for 12 h.

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Figure 8. Content of the two isoforms of phyA, phyA′ (light gray) and phyA′′ (dark gray), in % (a) and relative units (b) in coleoptile tips and mesocotyl base of etiolated maize shoots (LG 32.26 var.)—intact tissues and okadaic acid (0.5 μm) treated for 12 h.

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Table 1. Characteristics of phytochrome A in etiolated seedlings of maize (Kubanskaya var., if not indicated otherwise) and their changes upon treatment with specific PP1 and PP2A protein phosphatase inhibitors—okadaic and cantharidic acids
SampleTreatmentphy content, [Ptot], rel. unitsExtent of Pr to lumi-R conversion, γ1phyA′/phyA′′, %[phyA′]/[phyA′′], rel. units
Coleoptile tips (LG 32. 26)Intact tissues1.50 ± 0.340.35 ± 0.0170/301.05/0.45
Mesocotyl segments (LG 32.26)Intact tissues1.48 ± 0.220.28 ± 0.0558/420.83/0.65
Coleoptile tipsIntact tissues3.53 ± 0.200.32 ± 0.0264/362.26/1.27
Root tipsIntact tissues2.57 ± 0.210.31 ± 0.0362/381.59/0.98
Control in H2O
Coleoptile tips10 min H2O3.81 ± 0.040.30 ± 0.0260/402.29/1.52
16 h H2O2.78 ± 0.140.29 ± 0.0358/421.61/1.17
Mesocotyl segments (LG 32.26)16 h H2O1.29 ± 0.570.28 ± 0.0256/440.72/0.57
Root tips10 min H2O2.57 ± 0.210.31 ± 0.0362/381.59/0.98
4 h H2O2.40 ± 0.300.30 ± 0.0460/401.44/0.96
12 h H2O2.23 ± 0.120.29 ± 0.0458/421.29/0.94
Okadaic acid (0.5 μm solution)
Coleoptile tips (LG 32.26)12 h OA1.62 ± 0.570.16 ± 0.0332/680.52/1.10
Mesocotyl segments (LG 32.26)4 h OA + 12 h H2O1.28 ± 0.100.22 ± 0.0244/560.56/0.72
12 h OA0.78 ± 0.270.11 ± 0.0522/780.17/0.61
Coleoptile tips4 h OA2.50 ± 0.300.24 ± 0.0248/521.20/1.30
4 h OA + 12 h H2O2.50 ± 0.400.22 ± 0.0244/551.10/1.40
Mesocotyl segments4 h OA2.57 ± 0.190.24 ± 0.1849/511.26/1.31
Root tips10 min OA2.60 ± 0.400.31 ± 0.0363/371.70/0.90
4 h OA2.70 ± 0.200.30 ± 0.0460/401.70/1.00
4 h OA + 12 h H2O2.47 ± 0.150.31 ± 0.0162/381.53/1.00
Cantharidic acid (0.5 μm solution)
Coleoptile tips10 min CA3.90 ± 0.700.29 ± 0.0558/422.50/1.40
4 h CA3.06 ± 0.400.24 ± 0.0248/521.40/1.60
4 h CA + 12 h H2O2.95 ± 0.460.26 ± 0.0452/481.60/1.30
Root tips10 min CA2.50 ± 0.140.31 ± 0.0361/391.50/1.00
4 h CA2.58 ± 0.170.31 ± 0.0261/391.60/0.90
4 h CA + 12 h H2O2.41 ± 0.150.29 ± 0.0156/441.40/1.00

In case of mesocotyls, the spectra of the control and OA-treated samples are also practically identical (data not shown) and a 2.5-fold decline in the γ1 parameter is observed. However, Ptot is lowered by ca 40%. Thus, the most pronounced changes were observed with the γ1 parameter reflecting the redistribution in the content of the two phyA pools (see summary data in Table 1). It should be noted that, depending on the maize variety and samples used, there were two modes of γ1 decline—with and without decrease in the total phyA content, as in coleoptiles of the Kubanskaya var. and mesocotyl of the LG 32.26 var., on one hand, and coleoptiles of the LG 32.26 var., on the other, respectively. Cantharidic acid had no effect either on phyA in coleoptiles of maize upon the short-term (10 min) exposition, whereas a 4 h treatment produced a significant decline in the value of γ1 (Table 1). The following 12 h exposition of the 4 h-treated samples in water partially reversed the effect of the inhibitor. Ptot remained unchanged for the two treatments. As in the case of OA, the root tips were not sensitive to the CA action even after long-term exposure (4 h in CA + 12 h in water): the Ptot and γ1 values were comparable with those after the same time of incubation in water.

Evaluations of the phyA′/phyA′′ proportions show that in the control samples of coleoptile tips phyA′ dominates over phyA′′ (64/36% or 70/30% depending on the variety; Fig. 8, Table 1) and in the mesocotyls (var LG 32. 26) this domination is lower (58/42%; Fig. 8). However, upon okadaic and cantharidic acid treatment, a significant shift in the phyA′/phyA′′ equilibrium is observed: the phyA′/phyA′′ proportion is either almost entirely reversed or considerably shifted to phyA′′. Thus, we observed both redistribution of the concentrations between phyA′ and phyA′′ and decline in the total phyA content which is almost entirely achieved by the drop in the phyA′ amount (Table 1).

Interestingly, experiments with these specific phosphatase inhibitors on roots, contrary to our expectations based on the above experiments with coleoptiles and mesocotyls, did not produce any meaningful effects on the phyA parameters under all tested conditions (Table 1). This indicates that in roots treatment with okadaic and cantharidic acids has no influence on the different phyA species (Table 1).

Effect of animal and bacterial phosphatases on the phyA pools is lacking in crude extracts from etiolated maize and Arabidopsis seedlings

Analysis of the effects of the specific inhibitors of protein phosphatases shows that they modify the phyA′/phyA′′ equilibrium either by the phyA′′ pool increase at the expense of phyA′ (with Ptot remaining rather constant) and/or due to a preferential destruction of phyA′ (with a decline in Ptot). To verify a possibility of the direct phosphorylation/dephosphorylation of the phyA molecule as a source of its diversification into the two pools, we tried to observe a possible transformation of phyA′ into phyA′′ in crude extracts from maize and Arabidopsis upon addition of the animal or bacterial phosphatases active with regard to the serine/threonine residues—alkaline calf intestinal phosphatase (CIAP) and λ-protein phosphatase (Lpp). We have observed effects of (1) the type of buffer used (HEPES or Tris) on the properties and content of phyA, (2) Mn2+ on the yield of phyA fluorescence and (3) extraction procedure on the phyA′/phyA′′ ratio as compared with the intact maize tissues. However, there was no effect on the phyA′/phyA′′ proportion connected with the action of both phosphatases. No effect on phyA pools was observed on phyA in extracts from Arabidopsis as well upon treatment with CIAP: γ1 is virtually the same as in Arabidopsis in vivo (hypocotyls). This result might suggest that the assumed phyA phosphorylation as a means of the phyA′′ [RIGHTWARDS ARROW] phyA′ transformation is not true. However, it remains inconclusive because the lack of the effect in crude extracts could be due to the inhibition of the phosphatase activity by proteases or to their inactivity toward phytochrome.

pH as a factor regulating the phyA′/phyA″ equilibrium in the cell

In root tips stained with FDA, we observed a fluorescence rise along with the root development. In terms of pH changes, fast increase in pH (from 5.7 to 6.2) was seen from the moment of radicle protrusion up to 2 mm length (stage 1) followed by a decline to pH 5.8 at the length of the root of ca 5 mm (stage 2). Then, pH increased from 5.8 to 6.4 in the tips of the roots of 10–25 mm length (what corresponds to the stages 2–3; Fig. 9). In the literature, apoplastic pH was estimated to vary from 5.1 to 6.2 (see Ref. [57] and the literature cited therein). The character of maize root development also reveals three zones: up to 1.9 mm, alkaline; 2–5 mm, weakly acidic and 6.5–8 mm, acidic, and in general similar correlation between the root growth and surface pH was followed [57, 58]. These pH changes are in good agreement with the changes in the phyA pools′ ratio—fast increase in phyA′ at the first stage, then dominance of phyA′′ and finally, at the stage 3—again domination of phyA′ over phyA′′ (Fig. 5).


Figure 9. pH of maize root tips (1–1.5 mm; Kubanskaya var.) at different stages of root development (measured as a total length, L mm).

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In the experiments on crude extracts from maize seedlings, it was also found that a small change in the pH value of the medium brings about a considerable redistribution in the phyA′/phyA′′ ratio: at pH ≤ 6.8 the pigment was represented almost entirely by the phyA′′ form, whereas at pH ≥ 7.5 phyA′ reached considerable proportions (up to 40–50%). A more detailed investigations of the γ1 parameter reflecting phyA pools' ratio as a function of pH in maize extracts (using Tris buffer with pH in the range 6.5–8.5) revealed a bell-shaped dependence with the maximum at around 7.5–7.6 what corresponds to cytosol pH of a plant cell in a functionally active state (Fig. 10). At these optimal pH values, phyA retains its properties and a relatively high phyA′/phyA′′ ratio whereas at pH lower than 7.2, phyA becomes unstable. In crude extracts of different pH, incubated at 0°C, γ1 drops down during 20–30 min suggesting transformation of the labile phyA′ form into the relatively stable phyA′′ (Fig. 11).


Figure 10. Dependence of the γ1 parameter of phyA (open circles and polynomial fit 1) (a) and of the phyA′/phyA′′ ratio (b) on pH in crude extracts from maize coleoptiles (LG 32.26 var.).

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Figure 11. Variations in the γ1 value reflecting the phyA′/phyA′′ equilibrium with time of exposition at 0°C of crude extracts (from etiolated maize coleoptiles, LG 32.26 var.) with different initial pH values.

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To find out possible effects of pH on the phyA states in vivo similar to those observed in crude extracts (see above), root tips of 3-day-old maize seedlings were soaked in buffer solutions of different pH. phyA characteristics in roots remained unchanged after 2.5 h exposition in water (pH 6.4). However, at pH 4.9 we observed a steep drop of the γ1 parameter practically to 0, suggesting that phyA is represented entirely by the phyA′′ pool and that the decline of [phyA] especially pronounced at longer expositions (3 h) is achieved primarily by the phyA′ destruction (Fig. 12). At pH 8.0, phyA in root tips also revealed a decline of γ1 although not to such an extent as at low pH (γ1 dropped down two-fold) and thus a two-fold lowering of the relative phyA′ content. These data and also those obtained on maize coleoptiles (not shown) are thus in good qualitative agreement with the effects of pH changes on phyA state in crude extracts from maize roots.


Figure 12. Variations in the phyA′ and phyA′′ relative content in maize root tips (LG 32.26 var.) upon their soaking in buffer solutions at different pH (phyA′—gray, phyA′′—black).

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Heterologously expressed phyA belonging to the phyA′′ type is not affected by phosphatases and pH variations

The above picture of the effects of protein phosphatase inhibitors in vivo and of pH in crude extracts on the properties of phyA prompted us to investigate if they could be the consequence of direct protonation/deprotonation and/or phosphorylation/dephosphorylation of the phyA molecule. For this, we have carried out experiments on heterologously expressed and purified Arabidopsis phyA—full-length (phyA FL) and N-terminal sensor domain fragment (phyAΔ3)—expressed in Pichia pastoris and E. coli. A small fraction of phyA FL expressed in transgenic P. pastoris was already detected in the cell as a holoprotein without any exogenous chromophore addition. It had the low-temperature (77 K) emission maximum λmax at 680 nm suggesting that it incorporated endogenous PΦB whereas the pigment reconstituted in vitro with PCB had the emission maximum λmax at 668 nm and excitation λmax, at 650 nm. The fact that the fluorescence emission spectra of the pigment in the cell at 668 nm and 680 nm belong to phyA was proven by a full reversibility of the fluorescence intensity changes in the cycle of the Pr [LEFT RIGHT ARROW] Pfr phototransformation upon FR-R-FR illumination at 0°C (two cycles for phyA in the cells of P. pastoris were performed; Fig. 13a). Both the phyA-PΦB in the cell and phyA-PCB in vitro were shown to belong to the phyA′′ photochemical type, i.e. γ1 ≈ 0 for both of them (Fig. 13b). This is in line with the previous findings on phyA expressed in yeast [20]. Observation of phyA in P. pastoris cells with λmax = 680 nm characteristic of phyA in Arabidopsis, on the other hand, supports the data by Wu and Lagarias [59] suggesting that P. pastoris synthesizes PΦB. Interestingly, such a short-wavelength position of the phyA emission maximum (λmax = 682 nm) was also observed in protruding root tips containing primarily phyA′′ (at the beginning of the developmental stage 1, see above).


Figure 13. Raw fluorescence emission spectra (λe = 630 nm) of heterologously expressed phyA with endogenous chromophore in P. pastoris in vivo at 77 K. (a) The spectra labeled as F0, F2 and F3 were taken after actinic FR-R-FR illumination at room temperature, respectively. Two cycles of such phototransformations were performed on one and the same sample proving that the spectrum belongs to phyA. (b) Determination of γ1: the spectra F0 and F1 (after actinic R illumination at 85 K) were taken at 77 K. The fact that spectrum F0 is close to spectrum F1, that is, the γ1 value approaches 0, suggests that phyA in P. pastoris belongs to the phyA′′ type.

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The fact that the intact phyA in the P. pastoris cells (phyA-PΦB adduct) and extracted purified phyA from P. pastoris and E. coli in solution (phyA-PCB adduct) are represented only by the phyA′′ type may suggest that autophosphorylation of the pigment is not likely to be a potential mechanism of the phyA′ formation (assuming that these pigments possess kinase activity). On the other hand, it might be possible that some plant-specific factors lacking in the expression systems are necessary for such a mechanism to operate. To verify this, we investigated a possibility of phyA plant-specific modification by mixing Arabidopsis phyA expressed in P. pastoris and reconstituted with PCB in vitro with crude extract from etiolated Arabidopsis seedlings. It was expected that phyA′′ would be partially converted into phyA′, as hypothesized for the posttranslational modification of phyA′′ to form phyA′ in plants (see 'Introduction'). The results on the samples with added crude extract with (1) pellet, (2)ATP, (3) ATP + CaCl2, and (4) ATP + MgCl2 had the same parameters as that with only buffer added with γ1 ≤ 0.1, indicating that the native, undestroyed plant cell conditions are essential for the phyA′′ modification. Finally, contrary to the above observations on phyA in crude extracts, we have found that pH variations (from 6.5 to 8.5) do not affect the state of the purified heterologously expressed phyA-PCB from P. pastoris and E. coli, both for the full-length and truncated pigment. In these systems at all the pH in the tested range we found phyA only in the initial phyA′′ form.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The purpose of this work was to improve our understanding of the nature of the two phyA forms, mechanisms of their posttranslational modification and means of maintaining proper phyA′/phyA′′ balance in the cell. For this, we used etiolated maize seedlings and followed variations in the spectroscopic and photochemical parameters of phyA, which depend on plant organ/tissues and on the stage of its development. In agreement with our earlier findings of the phyA heterogeneity [2, 17-19], we can interpret them as a manifestation of the presence of the two phyA species in the cell—the longer wavelength, at low temperatures photochemically active species phyA′ and the shorter wavelength at low temperatures photochemically inactive phyA′′. Evaluations of their content in tissues (this work and [43]) have shown that it changes in the growing etiolated plant, even tissue specific, as observed by the phyA′/phyA′′ content in a developing maize root. These changes can be correlated with the stages of organ and tissue development.

In the present and our previous works, we found a dependence of phyA′ and phyA′′ on the total phyA content such that phyA′′ dominates at low Ptot and reveals saturation with growing Ptot, whereas phyA′ almost linearly depends on Ptot (Fig. 6). There is a critical Ptot (ca 1/7 of maximal Ptot values) where phyA′ and phyA′′ are at equilibrium with each other indicating that there is a limiting factor for phyA′′ accumulation. Close similarity of these dependences in roots and in coleoptiles (Fig. 6) indicates that they might be governed by universal factors and processes. Understanding of the nature of these factors is important for elucidation of the structural difference between the two phyA forms and mechanism of regulation of phyA pools' content and functions. Understanding of the characteristics of the subpools and mechanism of their formation could also explain the differentiation between cytoplasmic and nuclear phyA functions and the differentiation of these two subpopulations in the cell.

Considering that phyA′ and phyA′′ could be a soluble and membrane-/protein-associated and/or phosphorylated and dephosphorylated species, respectively (see reviews [2, 18, 19]), we investigated the connection between the phyA′/phyA′′ content with the acidic/alkaline and phosphatase/kinase equilibria in the cell. These factors were shown to govern regulation processes in the cell [37, 42, 57], in particular, they may modify the state and functional activity of phytochrome. To analyze possible phosphatase activity with respect to the origin of the different phyA pools, we used established structurally unrelated inhibitors of protein phosphatases of the PP1/PP2A classes, specific for Ser/Thr dephosphorylation—okadaic and cantharidic acids. In these experiments, special attention was given to control viability of the treated tissues to exclude possible effects of cell disintegration on the phyA pools (see the 'Materials and Methods' section). In particular, we used the inhibitors at the low concentration (0.5 μm)—the same or well below those usually taken in the PP inhibitory analysis. The condition of the treated cells was continuously controlled by light and fluorescence microscopy observation as well as by low-temperature fluorescence measurements. These methods did not reveal any signs of cell damage after 4–12 h OA or CA treatments which, on the contrary, were well pronounced in the case of the cell purposefully killed with alcohol or 100-fold higher concentrations of the inhibitors. In the case of prolonged (12 h) treatment of coleoptiles with OA, there was a slight loss of turgor observable. This, however could not have influenced the measurements because even heavy dehydration of maize and barley coleoptiles (up to 85% loss of the initial fresh weight) did not affect the phyA′/phyA′′ equilibrium [60]. The specific character of the PP inhibitors' effects in different organs and tissues (see below) can be also considered as a circumstantial evidence that they are not simply due to the destructive action of the inhibitors. These inhibitors were found to be active upon long-term expositions (hours) shifting the phyA′/phyA′′ ratio toward the latter at the expense of the phyA′ destruction and/or the phyA′ [RIGHTWARDS ARROW] phyA′′ transformation. The character and extent of the okadaic acid effects varied depending on the maize line used (Kubanskaya or LG 32.26), what may be connected, taking into account the high variability of the phyA′/phyA′′ content and its dependence on the physiological state of the plant (see above), with differences in growth and development of the respective plant lines. The effect was also tissue/organ dependent—it revealed itself with different extent in coleoptiles and mesocotyls, but was lacking in roots. The explanation of the phyA′ [RIGHTWARDS ARROW] phyA′′ transformation would be that phyA′′ is the phosphorylated species on Ser/Thr residues whereas phyA′ is the dephosphorylated one whose content (relative and absolute) increases upon the shift in the phosphatase/kinase equilibrium by phosphatase inhibition. However, this appears not to be the case for the following reasons. First, the fact that phyA′ converses into phyA′′ in solution upon treatment with animal phosphatase and that the major part of phyA in the cell, which by our estimations is represented by phyA′, is phosphorylated and a minor fraction (phyA′′) is dephosphorylated (see 'Introduction') suggests that phyA′ is the phosphorylated species. Second, in the experiments with purified heterologously expressed phyA, we failed to observe transformation of phyA′′ into phyA′ upon treatment of the pigment with the animal and bacterial phosphatases. At the same time, this experiment does not obviously exclude the reverse situation, when phyA′ is phosphorylated and phyA′′ is dephosphorylated. However, contrary to our expectations, we did not observe the phyA′′ into phyA′ transformation upon the action of the specific PP inhibitors in maize tissues. Indeed, proceeding from the fact that phosphorylated phyA undergoes a more rapid light-induced destruction [33], we may postulate that a similar process, although at much lower rate, may operate in darkness: inhibition of phosphatases increases the phyA lability (via its increased phosphorylation) and/or activity of the phyA-destruction system. Similarly, we assume that the shift in the activity of the phosphatase/kinase system toward the latter could facilitate, for the same reason, transformation of the more labile phosphorylated phyA′ into presumably dephosphorylated and more stable phyA′′. It should be noted again that the more stable and presumably dephosphorylated phyA′′ species is not affected by the inhibitors indicating the specificity of their action. This concept is supported by the fact that the inhibitors were inactive in roots: phyA here is more stable than in shoots [61]. Another explanation of the inactivity of the specific inhibitors in roots is that FyPP (type PP2A), in fact, the only PP known to interact with phytochrome, is absent in roots [31], whereas those present in roots—PAPP5 [32] and PAPP2C [62]—are not of the PP1 or PP2A types. It should be noted in support of this that the lack of the effect of the specific PP inhibitors (OA and CA) in roots cannot be attributed to their inability to enter the root cell. Okadaic acid was shown to increase microtubule dynamics in maize root tips suggesting that phosphorylation is partly responsible for maintaining its high rate [39]. Interestingly, in the context of the current discussion, light changed cortical microtubular array in roots. These PP inhibition experiments suggest that the phosphatase/kinase balance in the cell strongly contribute to the stability and regulation of the phyA′/phyA′′ system in the developing seedlings.

A second factor affecting the phyA state in the cell appears to be the intracellular pH. We have obtained three lines of evidence suggesting that the acidic/alkaline equilibrium in the cell may regulate the phyA′/phyA′′ balance and, thus, phyA function in general. First, we observed in crude extracts from etiolated maize seedlings a clear rise of the phyA′/phyA′′ ratio with increasing pH values. It reached its maximum at the pH of 7.4–7.5, equivalent to the cytosolic pH at the cell's functionally active state and declined then at higher pH values. The phyA′/phyA′′ ratio observed in the extract at pH 7.4 correlated also with that observed in intact cells. Second, a qualitatively similar observation of the phyA states was made in situ—coleoptiles and roots kept in buffers at different pH. These data suggest that the variation in the pH on the outside of the cell affects the state of the phyA pools in the cytoplasm, most likely via changing its pH. And finally, a third argument for the correlation between phyA′ and pH is the observation of similar dynamics of their changes in the intact developing roots. The pH values obtained with the use of pH-sensitive dye FDA reflect the situation in the cytoplasm where it resides according to [63-65]. In our experiments, pH in the tips of growing roots appeared to be relatively low and changed after their protrusion and rose from there on to reach almost neutral values along with root growth (Fig. 9). These pH changes correlated well with changes in the relative content of phyA′. The former are also similar to those observed in the literature [57, 58]. It should be noted that, as in the case of the action of the PP inhibitors, this effect is not connected with direct protonation/deprotonation of the phyA molecule, but is rather a consequence of a modification of phyA transformation associated molecule(s) or processes. This is suggested by the lack of the effect of pH on purified heterologously expressed phyA′′—pH changes in a wide range (6.0–8.5) failed to produce phyA′ from phyA′′. The data obtained in this study are first evidence of the connection of pH and phosphorylation status in vivo on phyA function. As both effects appear not to be direct, further investigation is needed to identify the specific protein partners of phyA to which this changes are targeted.

Our study also implies that a possible phyA phosphorylation (presumably in the phyA′′ form) producing phyA′ is rather a process of trans- but not autophosphorylation because all heterologously expressed Arabidopsis phyA (as purified full-length and truncated phyA-PCB in solution or as intact phyA-PΦB in transgenic P. pastoris cells) was found to be only in the phyA′′ form, in agreement with the data from [20]. Testing if the modification requires plant-specific factors to form phyA′ we aimed to produce the phyA′′[RIGHTWARDS ARROW]phyA′ transition of purified heterologously produced phyA′′ by addition of Arabidopsis crude extract from etiolated seedlings. However, the observed minor changes in the spectroscopic and photochemical parameters suggesting accumulation of the phyA′ pigment type were too small to be attributed to a plant factor supplied with the extract.

What is clear now, however, that both the acidic/alkaline and phosphatase/kinase status of the cell contribute to the regulation of the phyA′/phyA′′ equilibrium in the growing etiolated seedlings and, hence, of their functional activity. In particular, variations in the phyA′/phyA′′ equilibrium (without considerable changes in the total pigment content) upon inhibition of the phosphatase activity in the cell may mediate the effect of such a modification on the expression of light-inducible genes (see Refs. [37, 39]). This is suggested by the fact that phyA′ is the most likely phyA species implicated in the modulation of gene activity (of those involved in de-etiolation, including the active protochlorophyllide formation, see Ref. [66, 27, 67]) and that its content declines upon treatment with the PP inhibitors.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This work was supported by the Russian Foundation for Fundamental Investigations (grants nos. 05-04-49549, 08-04-01453 and 11-04-01732 to V. S.) and the Deutsche Forschungsgemeinschaft (DFG; grants no. 436 RUS 17/99/04 to P. G. and V. S. and 436 RUS 113/925/0-1 to J. Hughes and V. S., ZE485/2-1 to M.Z.). We are grateful to Prof. J. Hughes for his continued support, to Drs. F. Grolig, J. Malliet, A. Maisuradze, N. Matveeva, D. Liebsh and J. Rösler for their helpful and stimulating discussions and also to Mr. M. Göttig for growing maize seedlings.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
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