PHYSIOLOGIA PLANTARUM 119: 295–308. 2003 Printed in Denmark – all rights reserved
Copyright # Physiologia Plantarum 2003 ISSN 0031-9317
Spatial and temporal changes in multiple hormone groups during lateral bud release shortly following apex decapitation of chickpea (Cicer arietinum) seedlings Johanna C. Madera R. J. Neil Emeryb,* and Colin G N. Turnbullc a
Department of Botany, The University of Queensland, Brisbane, Queensland 4072, Australia Department of Biology, Trent University, Peterborough, Ontario, K9J 7B8, Canada c Department of Agricultural Sciences, Imperial College at Wye, University of London, Wye, Kent TN25 5AH, UK *Corresponding author, e-mail:
[email protected] b
Received 19 November 2002; revised 26 March 2003
Although the co-ordination of promotive root-sourced cytokinin (CK) and inhibitory shoot apex-sourced auxin (IAA) is central to all current models on lateral bud dormancy release, control by those hormones alone has appeared inadequate in many studies. Thus it was hypothesized that the IAA : CK model is the central control but that it must be considered within the relevant timeframe leading to lateral bud release and against a backdrop of interactions with other hormone groups. Therefore, IAA and a wide survey of cytokinins (CKs), were examined along with abscisic acid (ABA) and polyamines (PAs) in released buds, tissue surrounding buds and xylem sap at 1 and 4 h after apex removal, when lateral buds of chickpea are known to break dormancy.
Introduction The mechanisms by which plants achieve their characteristic shape have been of longstanding interest, however, they are far from being satisfactorily understood. There is general consensus on the concept of control by polarity. The poles of the plant, shoot and root tips, are thought to produce hormones and co-ordinate the processes shaping plant architecture via gradients of these hormones while integrating signals from other organs (Werner et al. 2001). A major focus involves auxin [i.e. indole-acetic acid (IAA)] and cytokinins (CKs) as key players in the concept of control by polarity in plants. Pioneering work with exogenous hormones (Sachs and
Three potential lateral bud growth inhibitors, IAA, ABA and cis-zeatin 9-riboside (ZR), declined sharply in the released buds and xylem following decapitation. This is in contrast to potential dormancy breaking CKs like trans-ZR and transzeantin 9-riboside 50 phosphate (ZRMP), which represented the strongest correlative changes by increasing 3.5-fold in xylem sap and 22-fold in buds. PAs had not changed significantly in buds or other tissues after 4 h, so they were not directly involved in the breaking of bud dormancy. Results from the xylem and surrounding tissues indicated that bud CK increases resulted from a combination synthesis in the bud and selective loading of CK nucleotides into the xylem from the root.
Thimann 1967) led to an apical dominance model in which auxin, originating from the shoot apex, represses bud break and root-supplied CKs promote axillary bud break. The ratio of the two hormones is assumed decisive for the fate of the bud, and whether it remains dormant or starts growing. Intense efforts to verify the applicability of this model to the regulation of apical dominance by endogenous hormones have not been rewarded with a complete understanding of the mechanisms involved, although a wealth of evidence conforms to the auxin : CK ratio theory of Sachs and Thimann (1967). However, a number of controversial reports and inconsistencies of the CK : IAA
Abbreviations – AA, amino acids; ABA, abscisic acid; CKs, cytokinins; IAA, indole-acetic acid; PAs, polyamines; ZR, zeatin 9-riboside; ZRMP, zeatin 9-riboside 50 phosphate; DHZR, dihydrozeatin 9-riboside; DHZRMP, dihydrozeatin 9-riboside 50 phosphate; iPA, isopentenyl adenosine; iPAP, isopentenyl adenosine 50 phosphate; iP9G, isopentenyl adenine 9-glucoside; ZOG, zeatin 0-glucoside; Z7G, zeatin 7-glucoside; Z9G, zeatin 9-glucoside. Physiol. Plant. 119, 2003
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ratio as indicator of lateral bud release has fuelled further research. It appears that the model in its simple form fits under certain defined physiological circumstances, but in other cases may have to allow for the full complexity of processes contributing to axillary bud release (i.e. Cline 1991). Consequently, auxin–CK relations in apical dominance should be considered against the physiological background once bud release occurs. With this in mind, the present study advanced investigation of lateral bud release in a three-fold approach. Firstly, endogenous levels of multiple hormone systems were analysed to give a wider signalling picture of those implicated in the process of previous studies. To account for situations for which lateral bud release could not be explained simply by IAA : CK interactions, studies have turned to, and implicated, other hormone systems. These could involve abscisic acid (ABA), which has been found closely correlated with inhibition of bud growth (Gocal et al. 1991, Emery et al. 1998b). However, ABA appears not to be directly involved in bud release as the correlative signal, but as an additional factor, which can itself be regulated by dominant organs (Tamas et al. 1979a, b, 1981), or auxin application after decapitation (Knox and Wareing 1984). Further, Kotov and Kotova (2000) described age-dependent physiological characteristics influencing lateral bud out-growth such as a diminishing CK response to decapitation coinciding with decreases in IAA and gibberellins. Likewise, PAs have long been implicated in phenomena relevant to lateral bud break including dormancy control in a range of tissue systems such as embryos (Be´ranger-Novat et al. 1994), apple flower buds (Wang and Faust 1994), apple buds (Wang et al. 1986), and Helianthus tuberosus tubers (Scoccianti et al. 1993). PAs are plausible candidates for long-distance signalling since their transport is nonpolar, and involves xylem, phloem and parenchyma cells (Masse et al. 1989, Rabiti et al. 1989, Bagni and Pistocchi 1991, Beraud et al. 1992, Feray et al. 1992). Furthermore, PAs are reported to participate in the regulation of apical dominance in shoots (Burtin et al. 1991, Geuns et al. 2001) and branching in roots (Ben-Hayyim et al. 1994). Intriguingly, both apparently key branching hormones, CKs and auxin, can stimulate PA-synthesis (Rastogi and Davies 1991). Secondly, careful attention was paid to the timing of events, whereby endogenous hormone measurements were taken in the critical time window preceding, and leading up to, lateral bud release. There remains a lack of data on endogenous hormone levels in buds at the specific time of dormancy breaking. Short-term spatial and temporal resolution of hormone contents in buds, vascular system and surrounding tissue would be a key to identifying so-called switches that initiate bud growth. Few studies have quantified multihormonal levels in buds with simultaneous measurements of other correlative factors or potential switches involved in lateral bud growth preceding, and leading up to, the actual release point. In lupine detailed studies on branching patterns revealed lateral branch development was controlled by 296
interactive effects of endogenous CKs, IAA and ABA rather than by any single hormone (Emery et al. 1998b). Nevertheless, even in that study, there was low resolution for correlating the exact timing of bud break to hormonal levels. Likewise, the sole report on PAs in buds during their release from dormancy sampled, at earliest, 1 day after apical dominance release (Geuns et al. 2001). To better elucidate such switches, the multihormonal approach was combined with a reliable way of inducing and tracking bud break. The chickpea model system, developed by Turnbull et al. (1997) and later modified by Mader et al. (2003) allowed the capture of the endogenous hormone status shortly before, and during, axillary bud break. Accordingly, the dominant shoot was decapitated and CKs, IAA, ABA and PAs were carefully monitored on a time-scale of 1–4 h following the decapitation. Thirdly, the forms and sources of the potential hormone signalling agents were investigated by taking measurements from several tissue types and xylem sap. Due to the complexity of CK synthesis and metabolism, an approach was used to survey as wide a range of CK forms as possible. So far, despite inconsistencies (i.e. Faiss et al. 1997) shoot apex-sourced auxin and rootsourced CKs regulating the development of organs elsewhere in the plant remain central to all models on the integration of whole plant architecture (McKenzie et al. 1998, Sussex and Kerk 2001). Yet it has been recognized that this model of simple polarity falls short of explaining a diverse range of branching patterns among species (Cline 1996, Emery et al. 1998b). The answer may lay in where the hormones are produced, how they are translocated, and what forms of the hormone systems are active in various tissues and transport fluids. Although significant quantities of CK can be produced in the shoot, it is still envisioned that, in many cases, CKs are produced primarily in the root and then transported in the xylem stream driven by foliar transpiration to integrate root and shoot development (Emery and Atkins 2002). Evidence that this assumption applies to apical dominance release in chickpea comes from our previous study (Mader et al. 2003). When a pulse of labelled trans-ZR was introduced into the xylem stream, it was shown that active CKs in the xylem were diverted to enrich the bud released from dormancy directly after the loss of the dominant apex. In the present study we investigated how those effects of decapitation on transport and metabolism, which were previously observed by use of [3H]-CK tracers (Mader et al. 2003), conform to the distribution patterns of endogenous CK in xylem sap, the responding bud and its surrounding tissues. Furthermore, it was of interest to determine which distinct changes in CK forms correlated with the initiation of bud growth and whether metabolic changes relating to bud break could be detected. Currently the specific CK forms that are most effective in promoting or otherwise regulating specific developmental processes are poorly understood. Thus, special care was taken to isolate and conserve CK nucleotides as accurately as possible and to Physiol. Plant. 119, 2003
separate cis- and trans-CK isomers. By doing so, the present study advances initial findings on CK increases in lateral buds (Turnbull et al. 1997) when measures to quantify nucleotides reliably and to separate cis-isomers had not been in place. Like most CK research, the focus had been selectively on the trans-isomers. In the past this was not cause for much concern on the basis that cisisomers have been suspected to be artifacts formed from RNA during extraction and were generally regarded to be inactive (Kaminek 1982, Tay et al. 1986). Recently, evidence has indicated that cis-isomers are not artifacts, they may frequently predominate the CK profile (Emery et al. 1998a, Martin et al. 2001, van Rhijn et al. 2001) and speculation that they might be biologically active (Morris 1997) emphasized the need to include cis-isomers of a more complete understanding of CK physiology. In summary, the current study investigated whether shortcomings of the classic CK : IAA model of apical dominance might be explained by looking at a more detailed physiological background. This included: using a model system that captures the critical timing leading up to lateral bud release, analysing multiple groups of hormones that have been implicated in bud dormancy, measuring a greater range of CKs than has been traditionally considered, and examining potential hormone source organs and translocation in the xylem stream.
Materials and methods Plant material A chickpea model system was used as previously described (Mader et al. 2003). Seedlings (Cicer arietinum L. cv. Bumper) were grown hydroponically until the three-node stage. Plants with one fully expanded leaf at node 2 and one expanding leaf at node 3 were decapitated 5 mm above node 1. Xylem sap and plant parts as depicted in Fig. 1 were collected at 0, 1 and 4 h after decapitation, snap frozen in liquid nitrogen and stored at 80 C. Xylem sap collection was performed with a humidified pressure chamber. Cotyledons were removed, cuts wiped with AMP solution (10 mg ml1 water) to clear phosphatase released from damaged cells and sealed with parafilm. Then the stem was cut just below node 1. The cut was cleaned with AMP solution and blotted dry. Excess water was removed from the roots with soft tissue taking care not to break the roots. Up to three plants were sealed together. The 0 h control was collected from intact plants. For the 1 and 4 h samples serial decapitation of lots of three plants was followed by serial collection after either 1 h or 4 h. Two fractions were collected, one at 0.35 MPa for 10 min followed by a second fraction at 0.70 MPa for 5 min. Both sap fractions were collected in separate tubes containing AMP solution (0.1 mmol per 100 ml sap collected from three lots of three plants) to reduce phosphatase-related losses. These subsample tubes were kept on ice during collection and then stored at 80 C. Physiol. Plant. 119, 2003
Fig. 1. Plant parts analysed from intact and decapitated Cicer arietinum plants at the three-node stage with one fully expanded true leaf.
Yields for both intact and decapitated plants were about double the transpiration rate of an intact seedling, around 1.2 and 2.7 ml min1 per plant in fractions 1 and 2, respectively. Subsamples were combined to give a minimum of 1 ml sap for each time-point sample. To standardize any unavoidable enzymatic contamination from wounded cells, care was taken to collect sap from the same number of plants for each time-point sample. Collecting intact and decapitated plants at the same flow rate was necessary to get sufficient sap in the short collection period necessary to capture time-points. Cytokinin quantification Sample preparation Tissue was freeze-dried and ground in a ball mill. Sample sizes were around 50 mg dry weight for buds, stipule and cut stump, and 100 mg or more for other tissues. Enzymatic degradation of phosphates was minimized by addition of AMP (1 mmol per sample). Samples were extracted and sonicated four times in 4 ml 65% cold (20 C) methanol and kept cold at all times. The four serial extracts were combined and dried in vacuo. In lieu of extraction, xylem sap samples were immediately diluted with 10 ml 0.1 M acetic acid. Internal standards were 5 ng of deuterated CKs (Z, ZR, DHZR, iP, iPA, iP9G, ZOG, Z7G, Z9G and if required ZRMP, DHZRMP and iPAP; 297
Apex Organics, Honiton, Devon, UK), 50 ng 13C IAA [13C6]-IAA (Cambridge Isotope Laboratories, UK) and 100 ng deuterated [2H6]ABA (CSIRO Plant Industry, Adelaide, Australia). These standards were added to sap samples after defrosting and to tissue samples at the start of the first extraction. Quantification of CK phosphates Several procedures were tested and found suitable. Large tissue samples were split and one aliquot was used to quantify the CK bases and nucleosides, and another to determine total CKs after treatment with phosphatase, giving nucleosides as the difference of the two. For total CK aliquots, AMP was omitted from the extraction procedure as described above. For samples of small tissue availability, for which it was hard to get material for analysis (in particular buds and stipules), sample splitting was not practical. A reliable way of separating the phosphate fraction was required. Cation exchange with propyl sulfonic acid (PRS) columns (Bond elute 12102094; Varian Inc., Palo Alto, CA, USA) was found reliable. The PRS column was attached in series to a syringe tip C18 cartridge (Waters 051910, Sep–Pak C18, Waters Australia, Rydalmere NSW, Australia). Both were conditioned with 10 ml 100% methanol followed by 10 ml of acetic acid (0.1 M). The extracts were re-suspended in 100 ml 100% methanol and sap samples diluted in 10 ml of acetic acid (0.1 M) were loaded onto the columns. A further 5 ml of acetic acid (0.1 M) were passed through and the C18 cartridge detached from the PRS column. The C18 cartridge retained CK nucleotides, IAA and ABA and these were eluted with 6 ml 70% methanol and dried in vacuo. Nucleotides were degraded to nucleosides by incubation with phosphatase (P-0280, pH 9, 5 units in 1 ml per sample; Sigma St. Louis, MO, USA) for 4 h at 37 C. Further purification was achieved by butanol partitioning. Samples were partitioned four times in an equal volume of water-saturated butanol. Samples were dried in vacuo and PRS/C18 combination followed as described above to separate CKs from IAA and ABA. Free base CKs and glucosides were eluted from the PRS column with 7.5 ml 2 N NH4OH in 50% methanol. The eluate was dried in vacuo (38 C), taken up in 1 ml (aqueous, pH 9), and partitioned with butanol. The butanol fraction was dried in vacuo and re-dissolved in 250 ml 100% methanol followed by 10 ml deionized water, and taken through a C18 step as described above. Analysis of CKs Separation by HPLC A method separating cis and trans isomers was devised, since cis-CKs can be expected to be a major component of the CK profile in some chickpea tissues (Emery et al. 1998a). Moreover, van Rhijn et al. (2001) have shown that rapid separation procedures frequently used in high sample throughput liquid chromatography tandem mass spectrometry (LCMS-MS), have caused co-elution of the 298
isomers and probably resulted in considerable overestimation of trans-isomers in addition to a failure of cis-CK identification. A C18 column (ZORBAX BonusRP, 5 mm, 2.1 50 mm; Hewlett Packard) was used with an Alltech (Alltech Associates Australia Pty Ltd., Sydney, Australia) C18 guard column at a flow of 0.3 ml min1. Solvent A was 10 mM ammonium acetate and solvent B acetonitrile: 10 mM ammonium acetate (9 : 1 (v/v). The solvent gradient was as follows: 0% B at 0 min, then to 30% B at 15 min, 100% B at 16 min, 100% B to 22 min, 0% B at 23 min and 0% B to 30 min. Quantification by LCMS-MS CK quantification by LCMS-MS was performed according to Prinsen et al. (1995) applying correction factors for isotopic impurities and linear calibration curves. The HPLC was followed in-line by an API 3000 triple quadrupole mass spectrometer (PE Biosystems, Thornhill, Canada) equipped with a pneumatically assisted electrospray as ion–spray interface. Positive ionization mode was used (ion spray potential 5500 V, orifice potential 35 V, ring potential 200 V, 30 eV collision energy). The transitions monitored were the same as those for Prinsen et al. (1995) with the following exceptions: [2H5]Z: 225/ 137; [2H5]ZR: 357/225; iPA: 336/204. Analysis of IAA, ABA, polyamines and amino acids The fraction containing IAA and ABA was processed and analysed by GC-MS according to Emery et al. (1998b). Polyamines were quantified by HPLC as described in Mader (1995). Amino acids were measured by HPLC as phenylthiocarbamyl (PTC)-derivatives. The derivatization was done according to Rosenlund (1990). For the HPLC analysis methods and column T-4 were selected from Vasantis and Molna´r-Perl 1999). The amino acids detected with this method were: aspartic and glutamic acid, asparagine, serine, glycine, hisitidine, threonine, alanine, arginine, proline, tyrosine, valine, methionine, cysteine, iso-leucine, leucine, phenylalanine and lysine. However, the main focus was placed on asparagine as the expected dominant amino acid.
Results Effects of decapitation on endogenous CKs The CKs consistently found in quantifiable amounts in all tissues were cis and trans-ZR, cis and trans-ZRMP, DHZR, DHZRMP, iPA and iPAP. Levels of other CKs were too low to be satisfactorily quantified in all samples. Thus data on them are not presented in detail. Buds In dormant buds of intact plants, cis-ZRMP far exceeded the concentrations of other CKs (Figs 2–5). cis-ZRMP was up to 28 times more concentrated than trans-ZRMP and cis-ZR eight times more than trans-ZR. Decapitation caused cis-ZR to decline to half of its original Physiol. Plant. 119, 2003
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Fig. 2. Effect of decapitation on trans-ZR (black bars) and transZRMP (grey bars) in the lateral bud, its surrounding organs and xylem sap of chickpea seedlings. Standard errors are based on three replicates.
concentration by 1 h, with a small recovery thereafter. Decapitation did not immediately increase the other CKs in buds. In fact, at 1 h, levels of iPA, iPAP, DHZRMP
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Xylem sap Sap was collected at the site of the released bud to give a picture of the CK profile arriving at the bud. Interpretation of sap CK concentrations after decapitation might seem complicated by the fact that sap from decapitated plants was collected at the same flow rate as sap from intact plants and not at the actual evapo-transpiration rate. In the given experimental conditions the transpiration rate measured in decapitated plants was reduced by 4.6 times compared to controls. However, according to established principles of ion transport, the amount of solute transported per unit time is largely independent of the rate of xylem flow whereby concentration increases with decreased flow and vice versa (Letham 1994). However, it is recognized that dilution of some 14
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and trans-ZR generally declined (Figs 2–5). This effect was most pronounced for iPA and iPAP. trans-ZRMP remained stable and DHZR increased only slightly. However, in contrast to the cis-CKs, by 4 h large decapitation-induced increases set in. The nucleotides, transZRMP and DHZRMP, increased the most (by 22 and 4.7 times, respectively). ZR and DHZR increased by around 3.7 times and isopentenyl-CKs (iPA and iPAP, Fig. 5) reached levels only slightly greater than double. trans-Z was higher than trans-ZR (2.3 pmol g1 FW) in dormant buds and showed a pattern after decapitation similar to that of cis-ZR; accordingly, at 1 h, trans-Z had decreased (0.8 pmol g1 FW) and recovered its original level by 4 h.
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Fig. 3. Effect of decapitation on cis-ZR (black bars) and cis-ZRMP (grey bars) in the lateral bud, its surrounding organs and xylem sap of chickpea seedlings. Standard errors are based on three replicates.
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Physiol. Plant. 119, 2003
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Fig. 4. Effect of decapitation on DHZR (black bars) and DHZRMP (grey bars) in the lateral bud, its surrounding organs and xylem sap of chickpea seedlings. Standard errors are based on three replicates.
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iPAP concentration (pmol g–1 FW or pmol ml–1) Fig. 5. Effect of decapitation on iPA (black bars) and iPAP (grey bars) in the lateral bud, its surrounding organs and xylem sap of chickpea seedlings. Standard errors are based on three replicates.
xylem compounds may not be proportional to flow (Beck and Wagner 1994, Else et al. 1995) and that there is considerable debate of xylem sampling methods (Bacon et al. 2002). Presently, because of difficulty in obtaining sufficient quantities of sap in decapitated plants, it is assumed that concentrations collected at a similar flow rate are directly comparable. In general, CKs increased in the xylem sap after decapitation. The cis and trans isomers of ZRMP and ZR comprised the main CKs (compare Figs 2 and 3 with Figs 4 and 5). In the first sap fraction collected at 0.35 MPa, trans-ZRMP had almost doubled in concentration and increased more markedly than trans-ZR by 1 h after decapitation. Thereafter, both these CKs kept increasing to about 3.5 times of their initial concentration. In intact plants, the cis-CKs were at least as abundant as trans-CKs (compare Figs 2 and 3). However, unlike trans-ZRMP, cis-ZRMP levels did not respond to decapitation. The cis-ZRMP remained relatively unchanged during the entire time-course (Fig. 3). cis-ZR was similar and even showed a small tendency to decline at 1 h. trans-Z was barely detectable in intact plants (0.03 pmol ml1) with only little change (to around 0.04 pmol ml1) after decapitation. Dihydro-CKs (Fig. 4) and isopentenyl-CKs (Fig. 5) were abundant enough for detection. DHZR and DHZRMP increased less than trans-ZR and trans-ZRMP at both time-points. Although iPA was unchanged at 1 h, it increased 4.5-fold by 4 h; iPAP, on the other hand, was reduced, to 300
30% and 75% of its initial value by 1 h and 4 h, respectively (see sap 0.35 MPa in Fig. 5. Note that sap collected at 0.7 MPa did not show this reduction, which will be discussed below.). Xylem sap results must be considered while bearing in mind the possibility that increased concentrations of sapCKs in decapitated plants were due to distribution being restricted by reduced transpirational surface, while biosynthesis remained unchanged. If this was the case, the sap CK concentration should increase. However, when the flow recommenced artificially with sap collection, the CK accumulation due to the ‘dead end’ effect would be flushed out in the first fraction. After that, in the second fraction, the concentration would decline once again, and approximate the situation in an intact plant. If, however, the concentration stayed high, this would indicate increased biosynthesis. The second fraction was collected at double the pressure, which resulted in a higher flow rate, and gave the opportunity to assess xylemloading characteristics of CKs. The cis and trans-ZR decreased in fraction 2 at all time-points in a manner approximately inversely proportional to the increased flow rate (Fig. 6). This is indicative of compounds undergoing little change in loading characteristics. On the contrary, nucleotides were increased compared to fraction 1 (Fig. 6). Since the 0 h controls had increased nucleotides, this was further evidence of greater xylem loading. Nevertheless, the dynamics of decapitationinduced effects on nucleotides was similar at both flow rates: little changes for cis-ZRMP and increases in other CK nucleotides, especially trans-ZRMP and DHZRMP. For iPAP, the increased loading at higher flow prevented its levels from falling after decapitation in the manner observed at lower flow. Sustained increases of CK in both xylem fractions strongly indicated CK biosynthesis was induced by decapitation.
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Fig. 6. Effect of increased flow rate, manipulated by externally applied pressure, on CK concentrations in xylem sap of chickpea seedlings. Physiol. Plant. 119, 2003
Stipule The stipule was the only transpiring leaf remaining once a plant was decapitated; therefore it was expected to attract and concentrate the most CKs (assuming CK were not selectively re-directed). In fact, the CK levels increased similarly to those of the buds (Fig. 2). However, concentrations per gram fresh weight, were about half of what was measured in buds. Using dry weights in place of fresh weights did not change this observation. cis-ZRMP levels (Fig. 3) did not change after decapitation so that the decline in the cis:trans ratio at 4 h (Fig. 7) resulted from increased trans-ZRMP levels only.
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Cut All CKs were increased at the cut stump (Figs 2–5). However, despite the concentrating effects of the delivery ‘dead end’ and the drying-out in the wound not all CKs increased to the extent one would expect. In the cut the accumulation pattern of individual CKs differed remarkably from that in the stem section (node) below. In particular, iPA and iPAP increased only four- and 10-fold compared with 37- and 66-fold in the node below over 4 h. With the exception of iPAP, nucleotide accumulation reached a plant-wide maximum in the ‘dead end’. trans-ZRMP increased 57-, DHZRMP eight- and cis-ZRMP 5.7-fold as opposed to respective 33-, eightand one-fold increases in the node below. In contrast to their nucleotides, trans-ZR and DHZR accumulated less in the cut versus the node below (trans-ZR two versus 41 times, DHZR two versus 11 times). The cis-ZR increased in a manner consistent with the drying out of the cut. This differential increase pattern of CKs could reflect differcut node internode cot node root mature root young
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2
ZR(MP) cis/trans ratio Fig. 7. Effect of decapitation on cis/trans ratios of ZRMP in lateral bud, their surrounding organs and xylem sap of chickpea seedlings. Physiol. Plant. 119, 2003
ences in susceptibility to degradation by CK oxidases or a selective redirection away from the cut. Plant patterns Considering the relative CK profile in the different plant parts (Table 1), it appears that the bud, stipule, young root, sap and cut, predominated by active CKs, were distinctly different from leafy tissue, stem and mature root, which were enriched in early pathway isopentenyl forms. Before cutting, there was a concentration gradient going down the plant, in the stem, the leafy organs, as well as in the roots (Figs 2–5). Leafy tissues representing the end of the transpiration stream were higher in CKs than stem tissue. The CK levels were highest in the shoot apex, followed by the expanding leaf (leaf 2) and the fully expanded leaf (leaf 1). In these organs trans-ZRMP (Fig. 2), iPAP (Fig. 5) and DHZRMP (Fig. 4) were present in similar concentrations. DHZRMP was low in other tissues. cis-ZR (Fig. 3) was generally higher than trans-ZR (Fig. 2). Upon decapitation CKs increased at different rates among the various tissues. At 1 h after decapitation, in mature roots and stem tissue, iPAP and DHZRMP started to increase. iPAP increased more than iPA, possibly indicating early steps of CK synthesis. trans-ZRMP had not changed along the plant by 1 h, except for a minor increase in free trans-ZR in the young root. At 4 h, concentrations of all CKs intensified in stem tissues with increasing distance from the root. This pattern was most noticeable for iPA and trans-ZR, whereas it was less evident for DHZR. Apart from the cut, the released bud was the only organ in which trans-ZRMP increased at a higher rate than trans-ZR. This is in contrast to iPAP (the precursor of ZRMP), which increased more than its nucleoside in all tissues except for the young root and xylem sap. Cis versus trans isomers of total ZRMP The cis-ZRMP was more abundant than trans-ZRMP in all parts of intact plants, including xylem sap and buds (Fig. 7). However, cis/trans ratios changed with decapitation. At 4 h after decapitation cis-ZRMP remained more abundant than trans-ZRMP only in roots. The cis and trans isomers were not affected by decapitation in the same way. Levels of cis-ZRMP did not change much in shoot or root tissue nor xylem sap in response to decapitation. A moderate increase was found only in the dried cut stump and the second fraction of root-sap (roughly two and 1.4 times, respectively). However, in buds, cis-ZRMP levels fell to less than half of those of intact plants within 1 h of decapitation. By contrast, trans-ZRMP responded to Table 1. Relative CK profile in different plant parts. Note: nucleotides and nucleosides were combined for this table. IPA5trans-ZR ¼ DHZR IPA ¼ trans-ZR . DHZR trans-ZR . DHZR . iPA IPA . trans-ZR . DHZR DHZR . trans-ZR . iPA trans-ZR . iPA . DHZR IPA . DHZR . trans-ZR
apex leaves bud, stipule, young root node, internode, cot node: stem root sap cut mature root
301
decapitation by increasing 13.5-fold in the buds largely between 1 and 4 h. The decrease in cis-ZRMP and increases in trans-ZRMP changed cis:trans ratios from 17 : 1 to 7 : 1 (by 1 h) and 1 : 2 (by 4 h) (Fig. 7). Furthermore, for ZROG it was noted that the cis-isomer was present in much higher quantities than the trans-isomer and, like cis-ZRMP, it was not influenced by decapitation.
Xylem IAA or ABA concentration (pmol ml–1)
1400
Effects of decapitation on endogenous IAA and ABA
cadaverine 25 20 15 10 5 0 putrescine 5
3 2 1 0
1 2 3 4 Hours after decapitation
Fig. 8. Effect of decapitation on IAA ( * ) and ABA ( * ) concentrations in lateral buds released from dormancy following apex removal in chickpea seedlings. Standard errors are based on three replicates.
bud
4
stipule
5
spermidine
stem/cut
6
0
302
30
stem
7
1 2 3 4 Hours after decapitation
node
8
200
(Fig. 10). Spermine was also present but only in consistently low quantities and was therefore not included in further analyses. Conjugated PAs data were not included
root
Bud IAA or ABA concentration (nmol g–1 FW)
9
400
Fig. 9. IAA and ABA in xylem sap collected at external pressures of 0.35 and 0.70 MPa following apex removal in chickpea seedlings. Standard errors are based on three replicates. *, IAA 0.35 MPa; &, IAA 0.70 MPa; *, ABA 0.35 MPa; &, ABA 0.70 MPa.
0 1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0
10
600
internode
PAs detected comprised the diamines cadaverine (CAD) and putrescine (PUT), and the triamine spermidine (SD)
800
0
cot node
Effects of decapitation on endogenous polyamines
1000
0
µmol g–1 FW
Following decapitation, bud IAA and ABA levels declined only moderately (Fig. 8). This same trend occurred in stem tissue and the stipule (data not presented). Overall, the spatial characteristics of decapitation-induced changes between ABA and IAA were distinctly different. IAA declined most at the decapitation site, whereas ABA increased at that site. ABA fell well below the control level of intact plants and this trend was more pronounced with distance along the stem, reaching its minimum at the shoot base. In roots, levels of both IAA and ABA were close to those of intact controls. IAA and ABA levels fell in xylem sap after decapitation (Fig. 9) in a response that was more pronounced for ABA. For both hormones, it was a greater decline than those observed in any of the tissues. The decline was quite rapid, for example, in the case of IAA, as early as 15 min after decapitation the level had decreased by 50–60% (data not shown in detail). Although sap collected at the higher flow rate decreased in hormone concentration, it did not occur proportionally to flow. For example as sap flow increased about 2.3 times, the ABA concentration decreased only 1.5 times, and remained uniform thereafter. IAA concentrations decreased similarly to those of ABA by 1 h, but after 4 h it reached the expected level that would result from dilution proportional to increased flow.
1200
Fig. 10. Effect of decapitation on free polyamines in the lateral buds and surrounding organs of chickpea seedlings. Standard errors are based on three replicates. 0 h control (white bars), 1 h (grey bars) and 4 h (black bars) after decapitation. Physiol. Plant. 119, 2003
for tissues, since they were usually undetected and always accounted for less than 10% of the total PAs. The tissue types differed in their profile of free PAs. For example, stem tissue had 87–90% CAD, 8–12% PUT and 0.7–1% SD. In roots PUT accounted for 20% and CAD for close to 80%. In contrast, in green plant organs (apex, expanding and expanded leaf) CAD was not necessarily the dominant PA, accounting for only 20–45%. PUT and SD contributed 15–50 and 10–40%, respectively. The quantitative data presented in Fig. 10 is restricted to plant organs that were below the decapitation site. A very similar PA-profile was observed in buds, stipules and adjacent nodes (Fig. 10). The relative distribution was not influenced by decapitation, although slight increases in all three PAs were seen in the stipule, the internode next to the decapitation site and the cut stump. Decapitation had only a relatively small effect on PAs in buds, where there was decreased CAD and SD. In xylem sap, the PA-profile differed markedly from that of tissues. Firstly, conjugated PAs were routinely detected in significant quantities. Conjugated PUT and CAD were at 0 h around 60 and 65%, respectively; at 0 h and at 4 h they were around 90% of the total PAs. Secondly, total PA levels were up to about 50 times more dilute than in tissue and concentrations of PUT and CAD decreased greatly with decapitation. Total PUT decreased to 50% by 4 h, and total CAD to about 25%. For the free PA fraction the decline was even more dramatic with reductions to 14 and 6% for PUT and CAD. Effects of decapitation on endogenous amino acids A wide range of AA was detected in sap and buds. In xylem sap asparagine was the dominant AA accounting for around 76% of the 16 AA measured (Table 2). All other AA were less than 4.5%. Decapitation did not significantly affect sap AA composition or concentration although there was a relatively small decrease in asparagine by 4 h. Furthermore, the dilution of AA in the second fraction at 0.70 MPa was proportional to the higher flow rate with no indication for increased xylem loading. Therefore, xylem AA was independent of decapitation and could be used as an internal reference for flow-dependent xylem loading of other sap components. This approach has previously been used to determine CK
delivery rates (Ma et al. 1998). Ratios of CK : AA (Table 3) support earlier results on stimulation of CK synthesis following decapitation and selective xylem loading of nucleotides. For example trans-ZR and cis-ZR delivery relative to AA did not change with increased flow rate, whereas the delivery of their nucleotides was raised. In buds the relative distribution of AA was different (data not presented) from that found in xylem sap. Asparagine was still the most dominant AA comprising approximately 15% of total AA in buds of intact plants. Other AA followed relatively closely in abundance and were approximately 10–14% or less. Decapitation resulted in increased bud AA, in particular of asparagine, which reached a relative abundance of 30 and 35% at 1 and 4 h, respectively. This represented an increase in asparagine concentration from 16 to 28 and 32 mmol g1 FW at 1 and 4 h, respectively.
Discussion Involvement of different hormone groups The present study assembled a picture of several hormones or hormone groups that are thought to play major roles in apical dominance release and plant architecture shortly following apical bud decapitation in the Cicer arietinum model system. Results are consistent with the classic hypothesis of a promotive effect of CK on lateral bud growth. However, it was emphasized that particular attention must be paid to CK form and function. The greatest change was a large increase in transZRMP, a CK thought to have strong promotive effects on the cell cycle (Riou-Khamlichi et al. 1999) and bud growth. Early, at 1 h after decapitation, xylem sap transZRMP increased dramatically and by 4 h, when bud growth was thought to have strongly commenced, trans-ZRMP levels not only remained high in the xylem sap, but also had dramatically increased in the lateral bud. Other shoot tissues accumulated large amounts of CK by this point. Consistent with our previous study (Mader et al. 2003) it appeared that the accumulation of trans-ZRMP in the released bud may have been mainly xylem translocated from the root and less from synthesis in the bud. In particular, trans-ZR increased in the xylem, whereas the released bud was the only tissue in which trans-ZRMP increased at a higher rate than transZR. This is significant since nucleotides have low
Table 2. Amino acid concentrations (mM) in root sap collected at 0.35 and 0.70 MPa from intact (0 h) and decapitated (1 h and 4 h) plants. Standard errors are in brackets (n ¼ 3) 0.35 MPa AA (mM)
0h
Asparagine Valine Glutamine Proline Aspartic acid Glutamic acid Arginine
17.57 0.90 0.61 0.38 0.36 0.20 0.05
Physiol. Plant. 119, 2003
0.70 MPa 1h
(±1.05) (±0.03) (±0.30) (±0.01) (±0.21) (±0.07) (±0.00)
17.61 0.95 0.57 0.40 0.37 0.20 0.05
4h (±0.33) (±0.09) (±0.24) (±0.04) (±0.23) (±0.09) (±0.00)
15.41 0.91 0.53 0.33 0.33 0.21 0.06
0h (±0.02) (±0.02) (±0.22) (±0.01) (±0.21) (±0.09) (±0.00)
9.45 0.54 0.49 0.21 0.16 0.08 0.03
1h (±1.05) (±0.07) (±0.19) (±0.03) (±0.09) (±0.04) (±0.00)
9.29 0.45 0.40 0.17 0.11 0.06 0.03
4h (±1.09) (±0.11) (±0.09) (±0.04) (±0.06) (±0.02) (±0.01)
8.51 0.49 0.46 0.15 0.15 0.07 0.04
(±1.51) (±0.07) (±0.15) (±0.01) (±0.10) (±0.03) (±0.00)
303
Table 3. Ratio of individual CKs (pmol ml1) and total AA (mmol ml1) in xylem sap collected at 0.35 and 0.70 MPa. trans-ZR:AA
0h 1h 4h
cis-ZR:AA
trans-ZRMP:AA
0.35 MPa
0.70 MPa
0.35 MPa
0.70 MPa
0.35 MPa
0.70 MPa
0.35 MPa
0.70 MPa
0.21 0.27 0.80
0.24 0.33 0.98
0.42 0.30 0.47
0.39 0.30 0.52
0.14 0.26 0.49
0.59 1.82 2.43
0.17 0.19 0.19
0.75 1.16 1.23
membrane permeability (Laloue et al. 1981, Laloue and Pethe 1982) and reduced susceptibility for CK oxidase (Chatfield and Armstrong 1986, Laloue and Fox 1989, Kaminek and Armstrong 1990), which would make trans-ZRMP a good form for sink tissue CK accumulation. By contrast, trans-ZR, has greater membrane permeability and is considered an efficient transport CK. Dihydro-CKs appeared to play a lesser role, and showed only modest increases in concentration. The cis-CKs responded quite differently to decapitation and their levels were either unresponsive or they frequently declined, especially in the released bud. IAA levels in released lateral buds and xylem sap declined as early as 1 h after decapitation and so this also fitted with the classic apical dominance hypothesis. However, the decline in bud IAA was more subtle, putting the emphasis on CKs as the apparent major correlative agents of bud growth. ABA also declined, which is consistent with the idea that it promotes dormancy of lateral buds (Emery et al. 1998b, Shimizu-Sato and Mori 2001, Sussex and Kerk 2001). While most confidence should be put into the IAA and ABA concentrations in the bud tissue, the present results also give an insight into the potential role of the xylem delivery of these hormones to the bud. It appeared that ABA and IAA varied with respect to xylem loading: ABA loading into the xylem increased consistently, but IAA only transiently increased. This is perhaps not surprising since theories of apical dominance often invoke basipetal polar auxin transport and imply symplastic movement rather than xylem transport (Tamas 1995). On the other hand, ABA is known to continually circulate through xylem and phloem (Wolf et al. 1990). Increased xylem loading in response to increased flow rate has been reported for ABA before (Else et al. 1995). Faster flowing sap may increase diffusion by steepening the diffusion gradient and raising the pH (Hartung et al. 1988, Sauter and Hartung 2002). Here the pH had increased from about 5.3 to 5.8. However, as with the situation for IAA, changes in ABA were dwarfed by changes in CK. Our results revealed no changes in polyamines in the early events leading towards lateral bud growth. In fact, the detected PAs actually declined following apical bud removal. They would have been expected to increase due to their involvement in cell elongation and division (Galston and Kaur-Sawhney 1995). Like Geuns et al. (2001) we found that PUT and SD levels may decline slightly but, unlike their results, no increase was observed in SPM, which was found at consistently minute levels throughout the experiment. Additionally, the present 304
cis-ZRMP:AA
study was the first to quantify CAD in the context of bud release, which did not vary significantly following apex removal. The sources and forms of CKs in released lateral buds The present data cannot establish, with certainty, whether decapitation stimulates CK synthesis in buds, roots or any other organs. Nevertheless, a general look at CKs among the various tissue and xylem sampled (Table 3) indicated that the bud, stipule, young root, sap and cut had profiles that would be expected for sink regions for CK in which active CKs should predominate. On the other hand, the leafy tissue, stem and mature root may be CK sources since they were enriched in early pathway isopentenyl-CKs. Following decapitation, increased CK concentrations and altered CK composition in both xylem sap fractions provided further support that stimulation of root CK synthesis occurred. This follows from the hypothesis that early CK biosynthesis occurs following nucleotide pathways (Astot et al. 2000, Kakimoto 2001, Haberer and Kieber 2002). Interestingly, the nucleotide proportion of total iPAP decreased after decapitation, from 50% in sap of intact plants to 25 and 15% in sap of decapitated plants over 1 and 4 h, respectively. This is logical considering iPA increased by 350% at 4 h, whereas iPAP fell by 70% by 1 h and was still decreased by 25% at 4 h. The reduction in iPAP probably also reflected an increase in its hydroxylation to trans-ZRMP. Results are consistent with this, considering that 1 h after decapitation trans-ZRMP increased in the first fraction of xylem sap by almost 95%, whereas trans-ZR increased only by about 30%. The second fraction of sap showed that pattern even more dramatically: trans-ZR levels had not changed, whereas trans-ZRMP increased by 150%. This also paralleled DHZRMP, which increased at a much greater rate than DHZR. The above observation that the probable ZRMPprecursor, iPAP, fell in concentration, while ZRMP itself increased, indicated that decapitation may promote CK activation (i.e. from an established pool of iPAP). However, since ZRMP can also be formed very rapidly from ZR (Mader et al. 2003) assessment is needed in the context of the whole network of CK pathways of biosynthesis, CK interconversion and catabolism. Fluxes through the other various branches of the pathways are affected by decapitation as well (Mader et al. 2003). For example, there seems to be a cut-specific metabolism, such as increased degradation and/or activation of other Physiol. Plant. 119, 2003
metabolic pathways triggered by elevated CK levels. The appearance of an unidentified compound in substantial amounts after the introduction of [3H]ZR into the xylem (Mader et al. 2003) is consistent with that possibility. Furthermore, the most apparent change in metabolism after decapitation was a reduction in CK catabolism. After 4 h, 90% of the [3H] label was in breakdown products versus less than 60% in buds of decapitated plants. This catabolic reduction was coincident with a decrease in auxin, which would reduce CK oxidase activity and any formation of nucleotides needed to resist oxidation. By 4 h, trans-ZRMP had increased relatively more than trans-ZR, shifting the ZR/ZRMP ratio from 1.4 to 0.44. Furthermore, Mader et al. (2003) observed significant changes in the CK metabolite profile. Intact plant buds contained more label as Z relative to ZR/ZRMP in comparison with decapitated plants. The endogenous Z concentrations measured here were in agreement with that observation. In intact buds, Z was equal to, or higher than, trans-ZR and, after decapitation, Z increased much less than ZR/ZRMP. A similar pattern was reported for buds of pea after decapitation (Blazkova et al. 1999). It needs to be verified whether this reflects a shift in CK metabolism, CK delivery routes, or both. The low Z-concentrations measured in xylem sap (in addition to the poor xylem connection of dormant buds (Sokorin and Thimann 1964) suggests that Z might be phloem-sourced. If Z was the main phloem transported CK and ZR the main xylem transported CK, this would explain the change in bud CK profile after decapitation. Isomers of unsaturated CKs In the present study it was apparent that cis and transZR levels were independently regulated. This agrees with others who suggest there is very little isomerization of cis to trans-CK (Wang et al. 1997, Suttle and Banowetz 2000). In buds, cis-ZR was halved within an hour of decapitation, whereas an increase of trans-ZR was slower to set in. This decrease in cis-ZR was unique to buds about to resume growth, which implied a possible involvement of high cis-ZR levels in promoting bud dormancy. It is consistent with an antagonistic role of cis-CK over trans-CK (Kuraishi et al. 1991), which could be down-regulated following apex decapitation, and a cis-CK : trans-CK ratio, which was halved by 1 h and declined sharply by 4 h after decapitation. Furthermore, at the cut site, trans-ZR did not accumulate to the levels found in adjacent stem sections, indicating a possible increase in CK oxidase activity. cis-ZR did not change in the same way, which is in line with cis-isomers being less susceptible to oxidases (Armstrong 1994, Bilyeu et al. 2001). These observations and the findings on trans- and cis-specific enzymes (Martin et al. 1989, 2001) reinforce the independence and precision of metabolic regulation. It signifies that cis-CKs likely play a more important part in CK homeostasis than previously acknowledged. Physiol. Plant. 119, 2003
cis-CKs have been neglected in the majority of CK research despite their predominance in many plants (Morris 1997, Emery et al. 1998a, van Rhijn et al. 2001), on grounds of their low biological activity in all classical CK bioassays (Kaminek 1982) and one more recent molecular assay (Haberer and Kieber 2002). However, one needs to bear in mind that bioassays are not representative for all developmental processes. For example in Mercurialis annua distinct trans-CK profiles are linked to male–female differentiation and the abundance of cis-CK correlates with sterility (Louis et al. 1990, Durand and Durand 1991). cis-CKs are found to be the predominant isomer in developing chickpea seeds (Emery et al. 1998a) and young lupine fruit (Atkins 1999). In fact, lupine fruit set and development correlates with distinct changes in the cis:trans ratio of CKs (Emery et al. 2000). In potato, cis-CKs were accredited to involvement of tuberization (Mauk and Langille 1978) and tuber dormancy (Suttle and Banowetz 2000). The possible role of correlative hormone interactions in the CK increase As discussed above, bud break and sustained growth rely on increased bud CKs as a combined result of changed (1) translocation, (2) synthesis in the root and the bud itself, and (3) metabolism. These factors are thought to be co-ordinated by correlative hormone interactions. Increased translocation of xylem CKs into buds after decapitation was previously demonstrated (Mader et al. 2003). The basis of this could be improved xylary strand connections similar to evidence in flax. There tracheary perforation differentiated as early as 1 h after decapitation and enhanced xylem flow correlated to early lateral bud growth (Raju and Marchuk 1993). An explanation can be derived from the auxin autotransport inhibition imposed by apical auxin at the junctions with the lateral buds (Bangerth et al. 1964). Once the inhibitory apex is removed basipetal auxin transport can commence in the bud. Consequently vessel differentiation can be expected to improve xylem supply. The CK increase might be self-reinforcing. CK are known to stimulate their biosynthesis (Meins and Hansen 1986). Moreover, forming a mutual interrelationship, root-CKs up-regulate auxins in lateral organs and shoot-auxins down-regulate CKs in roots (Bangerth et al. 2000). Further, auxin produced by the dominant apex autoinhibits its synthesis and export, and increases ABA levels in dominated buds. Here, the fall of bud IAA after decapitation can be indicative of bud auxin export following the loss of apically sourced IAA. The reduced auxin may help achieve the CK increase, not only by stimulating CK synthesis in the roots, but also by conserving CKs, since high auxin levels promote CK oxidase activity (Palni et al. 1988, Zhang et al. 1995). Regarding ABA, effects on CKs in bud break remain unknown. ABA has a diverse potential to modify CK metabolism (Mullins and Osborne 1970, Sondheimer and Tzou 1971, Back et al. 1972, Miernyk 1979, Whenham 1989, Tao 305
et al. 1991). The metabolic interactions between ABA and CK seem to operate both ways, since CKs enhance conversion of ABA to phaseic acid and stimulate overall ABA metabolism (Cowan et al. 1999). Furthermore, interconnections between ABA and IAA have been advocated, such as auxin inducting ABA formation (Tucker 1980), ABA affecting IAA uptake and transport (Pilet 1971) and ABA concentrations increasing as a direct result of auxin transport inhibition (Bangerth 2000). The increase in bud CKs triggers responses establishing dominance of the newly growing shoot. CKs generally up-regulate apical IAA synthesis (Bangerth et al. 2000) and, by affecting conjugation and decarboxylation of IAA, maintain an increased level of free IAA (Lau and Yang 1973) available for polar transport (Li and Bangerth 1992). Although this must be considered against evidence that transgenic plants overexpressing ipt indicated CK synthesis reduces IAA synthesis (Eklof et al. 2000). Furthermore, elevated CK levels in buds may stimulate auxin export out of the bud (Bangerth et al. 2000), thereby enhancing bud sink strength (Roitsch and Ehness 2000) and initiating greater CK biosynthesis in situ. Conclusions This study provided evidence for a sequence of changes, primarily involving CK form and quantity, in the induction of bud break following apex decapitation. Bud outgrowth involves cell division, cell expansion and DNAsynthesis. The sequence of these events depends at which stage of the cell cycle cells were arrested. In axillary buds of Cicer arietinum, mitosis was observed within an hour after decapitation (Hillman 1984), indicating cells were in G2. Commencement of the cell cycle this fast would need to be triggered by rapidly changing compounds. Three potential bud growth inhibitors, IAA, ABA and cis-ZR dropped in concentration and, were the first to be affected by decapitation. In fact, a decrease in IAA root sap concentration was seen as early as 15 min after decapitation. CKs, other than cis-ZR, started to show changes by 1 h after decapitation. CK levels accumulated shortly after and provided the requirements for DNAsynthesis. AA were seen to increase in buds within 1 h which suggests an involvement of AA in bud break and growth or, more likely, renewed sink strength. Furthermore, although PAs decreased in xylem sap quickly after decapitation, they did not change significantly in buds or other tissues after 4 h. This makes it unlikely that PAs are directly involved in the break of bud dormancy. It rather appears that their importance is linked with the process of subsequent growth but not with the actual breaking of dormancy preceding growth. A further key conclusion is that increases in CKs after decapitation are unlikely to be mainly a result of re-orientation of CK transport. Evidently metabolism and biosynthesis are regulated by way of interaction with other hormones to re-establish a shoot–root equilibrium after unbalancing disruptions in plant architecture. 306
Acknowledgements – We thank Dr Alun Jones for running the LCMS-MS samples, and Professor Dr Christa Critchley for making the work possible.
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