Further Adventures and Advances

The notion that pursuing higher education would preclude my returning to Hawaii proved wrong. A year before completing the Ph.D., I was offered and accepted a position in the newly formed Department of Food Science and Technology in the College of Tropical Agriculture at the University of Hawaii. By then I had two children, and a secure job was attractive. For the first few years, I pursued research related to agriculture and refrained from working on the xanthophyll cycle, expecting that another student in Chi's lab would take up the work. When it became clear that no one would, I returned to the xanthophyll cycle, focusing on the biochemistry with the long-range objective of gaining insights into function. During my xanthophyll cycle hiatus (1962-65), Achim Hager made significant progress on the cycle's biochemistry. He showed that violaxanthin de-epoxidase (VDE) was localized in the chloroplast lumen and required ascorbate and low pH for activity (Hager, 1966). The cycle's transmembrane organization (Fig. 3) was established when both groups showed that the reverse epoxidation of zeaxanthin to violaxanthin occurred on the stromal side of the thylakoid at near neutral pH in the presence of NADPH and O2 (Hager, 1975; Siefermann and Yamamoto, 1975).

Working on the xanthophyll cycle in Hawaii was not easy. Funding was limited, and there were no researchers nearby with whom I could interact that were engaged in related work on photosynthesis or carotenoid biosynthesis. Fortunately, grants from the

Fig. 2. 1974 photograph of David I. Sapozhnikov's group given to me by O. Koroleva when I visited St. Petersburg in 1989. Front row from left: I. Popova, D. I. Sapozhnikov, S. Eidelmann, O. Popova. Back row from left: E. Morkovskaja, M. Gabr, O. Koroleva, T. G. Maslova, and G. Kornjushenko.

Stroma Thylakoid Lumen

Fig. 3. VAZ transmembrane pathway for the xanthophyll cycle in higher plants.


Fig. 3. VAZ transmembrane pathway for the xanthophyll cycle in higher plants.

Fig. 4. Autographed banquet menu from the 1965 NATO Advanced Study Institute on the Biochemistry of Chloroplasts in Aberystwyth, Wales. In addition to names already mentioned, signatures by Trevor Goodwin, Giorgio Forti, Martin Gibbs, Norman Krinksy, Harold Strain, Jack Pries and Joseph Bradbeer, among others, are also present. How many signatures can you, the reader, recognize? It was exciting for me to be at this meeting of such notable scientists, most of whom I met for the first time.

Fig. 4. Autographed banquet menu from the 1965 NATO Advanced Study Institute on the Biochemistry of Chloroplasts in Aberystwyth, Wales. In addition to names already mentioned, signatures by Trevor Goodwin, Giorgio Forti, Martin Gibbs, Norman Krinksy, Harold Strain, Jack Pries and Joseph Bradbeer, among others, are also present. How many signatures can you, the reader, recognize? It was exciting for me to be at this meeting of such notable scientists, most of whom I met for the first time.

National Science Foundation, the U.S.D.A. Competitive Grants Program, and the Department of Energy allowed me to continue research on the VAZ cycle. These grants also enabled me to travel about once a year to a major meeting on photosynthesis. Given my isolation from the mainstream of photosynthesis research, the importance of attending these meetings cannot be overemphasized. The first international meeting I was privileged to attend was the Advance Study Institute on the Biochemistry of Chloroplasts held in Aberystwyth, Wales in 1965, sponsored by the North Atlantic Treaty Organization (NATO). I believe my invitation to attend came from Trevor W. Goodwin. While looking through memorabilia in preparation for this perspective, I found the menu that I had passed around for signatures at the farewell dinner meeting (Fig. 4). I hope readers can make out the names in this marvelous collection of signatures. Among them are Robin Hill, Tony San

Pietro, C. Stacy French, Dan Arnon, and many more, with apologies to those I have not mentioned.

Contact with the photosynthesis and plant biochemistry community has been an essential part of my forty-year stroll through the xanthophyll cycle and has created opportunities I might otherwise have missed. For example, in 1968 I spent my first sabbatical with Leo Vernon at the C.F. Kettering Research Laboratory in Yellow Springs, Ohio. There I met Teruo Ogawa, who was completing a postdoctorate with Leo, and with whom I became close personal friends. Teruo introduced me to the "opal glass" spectrophotometric technique perfected by Kazuo Shibata for measurement of light-scattering samples (Shibata, 1973). Upon returning to Hawaii, I applied the technique to chloroplast suspensions and found that violaxanthin de-epoxidation was detectable as a difference spectrum, with a peak at 505 nm, and could also be followed kinetically at 505 minus 540 nm (Yamamoto et al., 1972). This sensitive and rapid method for in situ measurement of xanthophyll cycle activity in chloroplasts was key for much of the progress we made during the 30 years that followed. Early applications of the spectrophotometirc assay included the discovery of the "availability" phenomenon, the intensity-dependent fractional release of violaxanthin from the total pool (Siefermann and Yamamoto, 1974); inhibition of VDE by dithiothreitol (Yamamoto and Kamite, 1972); and epoxidation of zeaxanthin to violaxanthin (Siefermann and Yamamoto, 1975). The method was also well suited for in vitro studies that demonstrated the requirement of lipid for de-epoxidation of pure violaxanthin (Yamamoto et al., 1974) and the substrate stereospecificity of VDE (Yamamoto and Higashi, 1978). The spectrophotometric assay of VDE activity remains useful to this day. It was recently applied to demonstrate that monogalactosyldiacylglycerol (MGDG), the major thylakoid membrane lipid, has a limited capacity to accommodate zeaxanthin and when this capacity is exceeded, stereospecific product feedback inhibition of VDE results (Hieber et al., 2004).

The serendipitous discovery of MGDG as the optimal chloroplast lipid for in vitro de-epoxidation of violaxanthin proved important. While I was able to obtain de-epoxidation of violaxanthin bound in washed thylakoid membranes, the same crude VDE preparation had no activity against purified violaxanthin, as had been reported by Hager (1966). Violaxanthin is insoluble in aqueous buffer, and various attempts to suspend or solubilize violaxanthin in a form that yielded activity failed. Isomerization and decomposition of the preparation were excluded as possible reasons. In the course of these tests, I ran out of the vio-laxanthin preparation that I had been using and, as a matter of convenience, recovered violaxanthin from "fat plates"* that Dorothea Siefermann, then a postdoctoral researcher in my laboratory, happened to be using for analysis of chloroplast pigments. Violaxan-thin that was eluted from these plates with acetone and used without further purification gave rapid and nearly complete conversion to zeaxanthin. The reason for this success was traced not to coconut oil from the plates but rather to a lipid component in the unsaponi-fied extract that co-chromatographed with violaxanthin. C. Freeman Allen earlier had separated the lipids in chloroplasts (Allen et al., 1966) and he kindly sent me samples that he still had on hand. All of the lipid samples we received supported de-epoxidation to varying degrees. We subsequently prepared a complete set of the major chloroplast lipids and found that MGDG was the most effective, giving rapid and complete de-epoxidation of violaxanthin in about 5 minutes under optimal conditions (Yamamoto et al., 1974). These results helped define the in vivo substrate of VDE: the violaxanthin that is converted to zeaxanthin is free in the membrane lipid phase rather than bound to pigment proteins. Exchanges between protein-bound pigments and free pigments in the lipid phase are implied. Recently, model systems consisting of soybean phosphatidylcholine only (Grotz et al., 1999) or egg phosphatidylchloline combined with MGDG (Latowski et al., 2002) have confirmed that lipid is required for "activation" of pure violaxanthin. However, de-epoxidation in these presumably bilayer systems were relatively slow and incomplete compared to de-epoxidation in the MGDG micelle system. MGDG constitutes a much larger fraction of the total chloro-plast lipid: 60% to phosphatidylcholine's 2% or less (Webb and Green, 1991) and thus the micelle system may more closely approximate the in situ environment of free violaxanthin. Whatever model system is employed, violaxanthin should be prepared from saponified extracts to avoid artifacts from even trace amounts of contaminating chloroplast lipid.

We used the pH-dependent binding of VDE to the thylakoid membrane and to MGDG to obtain the partial C-terminal sequence (Rockholm and Yamamoto, 1996), which was then used to clone the gene and express the VDE protein (Bugos and Yamamoto, 1996). The complete sequence showed that VDE was a lipocalin enzyme, the first identified in plants** (Bugos et al., 1998; Yamamoto et al., 1999). This finding confirmed conclusions drawn 20 years earlier-before lipocalins were known to exist-that the shape of the VDE active center resembled a deep well (Yamamoto and Higashi, 1978). The cloned VDE carried out the forward VAZ reaction, providing strong evidence that the reaction could be catalyzed by a single enzyme with mono-de-epoxidase function. Evidence that a single gene product accounted for de-epoxidation was shown by Niyogi et al. (1998), in which a deletion mutation in Arabidopsis inhibited all de-epoxidase activity. The cysteine rich domain in the N-terminal sequence and highly charged domain in the C-terminal sequence explained, respectively, the DTT inhibition (Yamamoto and Kamite, 1973) and the pH-dependent membrane binding of VDE (Rockholm and Yamamoto, 1996).

My walk through the xanthophyll cycle took several administrative detours from 1980-82, 1982-86, and 1994-96 as Acting Associate Dean of Research, Chair of the Department of Plant Molecular Physiology, and Director of the Hawaii Agricultural Experiment Station, respectively. During the second of these, another chance occurrence caused me to refocus on research. It was popular for a time to hold small, informal bi-national conferences in Hawaii. One such conference, on photoinhibition, was held in Honolulu in 1985. At that time the subject was outside of my field of interest, but I attended on invitation from David Fork, whom I knew from visits to Carnegie during my days as a graduate student. One report by Olle Bjorkman caught my attention. He showed the kinetics of chlorophyll fluorescence quenching resulting from photoinhibition, which I recognized as being similar to the kinetics of violaxanthin de-epoxidation. After the meeting, I wrote a research proposal, including a request for a pulse-amplitude modulated fluorom-eter (PAM)*** that I would need to investigate the possible connection between photoinhibition and zea-xanthin formation. The grant proposal was successful but I was "scooped" by publication of a seminal paper by Demmig et al. (1987) that reported the correlation between non-photochemical quenching (NPQ) and zeaxanthin formation. Barbara Demmig had, in fact, noted the possible correlation a few years earlier but had difficulty convincing others of its reality. (For an interesting account of the events surrounding her important discovery, see Demmig-Adams, 2003.) Later, Adam Gilmore showed by a modeling technique that antheraxanthin also contributed to NPQ as effectively as zeaxanthin (Gilmore and Yamamoto, 1993). It is now common practice to express de-epoxidation as the de-epoxidation state (DES), or (Z + A)/(V + A + Z), in conjunction with NPQ. The question of whether the correlation is a direct or indirect effect was recently answered with evidence that zeaxanthin is a direct quencher of excess energy (Ma et al., 2003).

Advances in research often result from the coupling of new analytical instrumentation or methods with the efforts of talented and dedicated individuals. This is certainly the case for contributions my laboratory made regarding the xanthophyll cycle and its relationship to NPQ. In terms of technology, the 505-nm absorbance change associated with de-epoxidation, the MGDG model system, and the HPLC method for resolution of zeaxanthin and lutein made significant differences in our research. The 505-nm change and development of the PAM provided an exceptional opportunity to examine xanthophyll cycle activity and NPQ simultaneously in chloroplasts. This application made it possible to show that although de-epoxidation and NPQ were both induced by light-dependent low pH, the protons for each were localized in different domains of the membrane (Mohanty and Yamamoto, 1996). I have not understood why the relatively simple opal-glass technique for the 505-nm change has not found more use, especially since it can be used simultaneously with NPQ

measurement. In contrast, the HPLC method we developed for analysis of plant pigments is in wide use today (Gilmore and Yamamoto, 1991). As with the identification of the 505-nm change and the development of the MGDG model system, we arrived at this method somewhat by circumstance. Thayer and Bjorkman (1990) had reported an HPLC method that separated lutein and zeaxanthin but the column they used was no longer available. Based on my previous experience in separating lutein and zeaxanthin by sequential partitioning and absorption columns, we looked for a column that had both of these properties. ODS-1 was identified as a possibility because of its light carbon loading and non-endcapping of active silyl groups. The column performed as we hoped.

Although the mechanism of quenching has largely been resolved, numerous questions about the xanthophyll cycle remain. The physiology of the cycle is not well understood. The pool size of violaxanthin and the fraction of the pool that is active in the cycle vary among plant species and growth conditions (Demmig-Adams et al., 1999, this volume). There is a growing body of evidence that the cycle's operations may be related to more than just NPQ (Yokthongwattana and Melis, this volume). If the cycle has multiple functions, how are these functions regulated? Mutant studies suggest that the cycle in not essential for photosynthesis (Jung and Niyogi, this volume) and yet, as far as I am aware, all wild-type plants have the xanthophyll cycle. Why has nature retained this complex, apparently multifunctional system if it is not of some critical advantage? Did the system provide the adaptability to light environments needed for terrestrialization over multiple generations? Is it simply coincidental that the dominant photosynthetic life forms in the ocean and on land have xanthophyll cycles, the diadinoxanthin cycle and the VAZ cycle, respectively? The xanthophyll cycle has been related to photoprotection of plants against sudden and prolonged light stress (Verhoeven et al., 2001) and to improved plant fitness, as indicated by seed production, under fluctuating light intensities of the natural environment (Kulheim et al., 2002). Interestingly, only half of the VAZ cycle (to antheraxan-thin) appears to be present in a few Rhodophyceae (Aihara and Yamamoto, 1968) and in Mantoniella (Goss et al., 1998). Are these species less fit? We recently proposed that zeaxanthin functions as a messenger in a signal-transduction network that operates in the lipid phase of the chloroplast membrane to explain the cycle's multifunctional capabilities (Hieber et al., 2004). As one who has been involved with the xantho-phyll cycle for nearly 45 years, I am surprised at how the questions seem never to end.

Fig. 5. Millie and I seated for lunch in Bagan during a recent tour of Myanmar.

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