Research Faculty #27
B.Sc. (1984), McGill Univ.;
Ph.D. Botany (1989) UBC;
Postdoctoral Fellow, (1993-95), Univ. Colorado
Research Associate, (1996-2000) UBC Vancouver
Assistant Professor (2000-2006) UBC Vancouver
Associate Professor (2006-2011) UBC Vancouver
Professor (2011-present) UBC Vancouver
Head of Botany Department, (2011-2016)
Visiting Professor, Nara Institute of Science and Technology, 2011
Academic Director of the UBC Bioimaging Facility (BIF) (www.bioimaging.ubc.ca).
In 2019-2020, this campus-wide Facility served 201 graduate student, post-doc, undergraduate, and faculty researchers from 108 research groups at UBC (87 groups), as well as other universities and institutions (8 groups) and industry clients (13 groups). A total of 180 UBC researchers (17 PIs, 49 Postdoctoral fellows, 72 PhD student, 15 Masters student, 14 undergrads, 8 staff, 5 visiting scientists) came from across 22 Departments in 8 Faculties.
BIF provides both service and training in confocal microscopy, scanning electron microscopy, transmission electron microscopy, electron tomography, cryo-fixation of cells, and cryo-TEM.
Plant cell biology, cellular basis of secretion of plant cell wall components; lignification in xylem development; ABC transporters and cuticle secretion
The Samuels lab studies how plant cells secrete their cell walls, both the polysaccharides and specialized cell wall components such as lipids and lignin. Our approach is to integrate cell biology with molecular biology and biochemistry to put cell wall biosynthesis and secretion into a cellular context. All plant growth, including agricultural and forestry production, is based on the organized assembly of plant cells into tissues, organs and whole plants. The plant cell wall determines the shape of the cell and connects cells into tissues and higher order structures, thus plant growth depends on cell wall production. In addition, terrestrial plants have evolved specialized regions of cell walls, such as the plant cuticle and lignified cell walls that are essential for water retention and water conduction, respectively. Lignified cell walls, such as those found in vascular tissues like wood, make the wall strong and waterproof. The removal of lignin from the cellulose of the cell wall has been identified as a barrier to enzymatic degradation of cellulose feedstock for biofuels, so there is strong interest in understanding lignified secondary cell walls.
Cellulose biosynthesis in secondary cell walls.
The formation of wood in the annual growth rings of trees is the most vivid example of how plant cells use carbon captured during photosynthesis to produce biomass that is rich in cellulose and lignin. Cellulose forms paper, textiles, building products, as well as having potential for the production of biofuels. Yoichiro Watanabe, a Botany Ph.D. candidate in my lab, co-supervised by Shawn Mansfield in the Faculty of Forestry, used live-cell imaging to quantitatively analyze cellulose synthase enzymes in developing xylem cells (Watanabe et al., Science 2015). This paper was featured in an accompanying ‘Perspectives’ article in Science. This was the first demonstration of live xylem cells actively depositing cellulose, and it has opened new areas of cellulose biosynthesis and bioenergy research.
In collaboration with Staffan Persson’s lab at Uni. of Melbourne, Yoichiro went on to discover how cells remodel their cellulose synthesis machinery during the switch from primary cell walls during their growth phase, to secondary cell walls during xylem development (Watanabe/Schneider et al. 2018 PNAS USA). One of the most exciting aspects of this work was the opportunity to observe both primary and secondary cellulose synthase enzymes in the same place at the same time. This work was featured in a commentary “Two types of cellulose synthesis complex knit the plant cell wall together” (Haigler, 2018 PNAS USA), which states “Watanabe et al. have opened the door to many research avenues that will allow us to fully explain and beneficially manipulate the synthesis of abundant renewable biomaterials.”
2. Lignin biosynthesis in secondary cell walls.
In addition to cellulose, lignin represents a significant (15-30%) component of the secondary cell wall mass of woody tissues. While lignin is important to plants' physiology by making the walls in the vascular system strong and waterproof, it makes break-down of plant biomass for renewable energy more difficult. Lignin forms when precursors called monolignols are crosslinked in the cell wall. Xylem vessels continue to lignify after they undergo programmed cell death, suggesting surrounding cells contribute monolignols for lignification. This “good neighbour” model was the subject of Rebecca Smith’s Ph.D. dissertation. We used a targeted gene knock-down to decrease monolignol biosynthesis only in lignifying cells (R. Smith and M. Schuetz et al., 2013 Plant Cell). Surprisingly, we found whether a lignifying cell had good neighbours or not depended on the context. In vascular bundles, good neighbours played an important role but in supportive fibers outside the vascular bundles, there were no good neighbours. By specifically manipulating lignin production in different cell populations, we could reduce or modify the lignin content of plants without affecting yield or vascular function (SmithPhD et al., 2015, 2017 Plant Physiol.). The finding that different populations of vessels and fibers cells have different mechanisms for wall lignification has interesting impacts for biotechnology, as it identifies different lignin pools that can be manipulated independently. Previous attempts to manipulate lignin for bioenergy crops used constitutive promoters that would alter lignin in all cell types, which has negative consequences for plant defense responses and growth. Here, we produced plants with improved release of cell wall sugars (saccharification) for biofuel production, yet the plants had normal vascular integrity, and biomass (Smith et al., 2017 Plant Physiol.).
Once monolignols are secreted into the secondary cell wall, they must be oxidized by enzymes called laccases and peroxidases, producing monolignol radicals that spontaneously cross-link into the lignin polymer. It is clear that lignin deposition is precisely spatially controlled in plant cell walls, and we discovered that specific laccases in Arabidopsis were responsible for directing lignin deposition (Schuetz et al., 2014 Plant Physiol.; Chou/Schuetz et al., 2018, J. Exp. Bot.). The impact of this work was to demonstrate that it is the activating oxidases, not the cell wall environment or other factors, that controls the spatial distribution of lignin in cell walls.
3. Characterization of glandular trichomes of cannabis.
The flowers of Cannabis sativa L. (cannabis) are used medicinally and recreationally by humans because they contain specialized cannabinoid metabolites. The metabolites are produced in hairs (glandular trichomes) on the flower. Early studies of cannabis had described three types of glandular trichomes (bulbous, sessile, and stalked), but their properties and relative contributions to producing metabolites were not known. We are studying the stalked glandular trichomes that dominate the cannabis flower. The cannabis stalked glandular trichomes have a large ‘head’ on a multicellular stalk. With advanced microscopy techniques, we can probe the interior of the stalked trichome head, demonstrating a proliferation of secretory cells and an extracellular storage cavity full of blue autofluorescent metabolites. This correlated with high cannabinoid content and enrichment of monoterpenes. By isolating glandular trichomes, we studied which genes are expressed during metabolite production. In addition to validating the proposed cannabinoid biosynthetic pathway, this transcriptomic data identified two new monoterpene synthases, and a suite of highly-expressed genes whose functions are presently unknown. Discovery of the unique properties of cannabis stalked trichomes, and the highly-expressed genes within them, is critical information for molecular breeding, targeted engineering, and optimized harvest and processing of this important plant.
4. Discovery of ATP-binding cassette (ABC) transporters in plant cell wall lipid export.
All land plants' aerial surfaces are coated with a lipid-rich waxy cuticle. The plant cuticle plays critical roles in plant ecophysiology by sealing the plant surface to prevent water loss. My research group was the first to discover that ABC transporters are required for lipid export from the epidermal cells to the plant cuticle (Pighin et al. 2004 Science). In collaboration with Carl Douglas, I co-supervised T. Quilichini, who extended this work by demonstrating that an ABC transporter is required for pollen cell walls formation in developing anthers (Quilichini. et al., 2010 Plant Physiol; Quilichini. et al., 2014 Plant Cell). This work used two-photon imaging of intrinsic fluorescence in anthers and cryo-fixation of anthers for electron microscopy (Quilichini et al., 2015 Annals of Botany). These ABC transporter studies are important because they demonstrated novel mechanisms of lipid transport to the cell wall.
Mendel Perkins, Ph.D. candidate
Sam Livingston, Ph.D. candidate
Jan Xue, M.Sc. student
Kenzie Arnott, M.Sc. student
Samuel King, Honours student
Hoffmann, N., Benske, A., Betz, H., Schuetz, M., Samuels, A.L. (2020) Laccases and peroxidases co-localize in lignified secondary cell walls throughout stem development. Plant Physiology DOI: https://doi.org/10.1104/pp.20.00473
Sutthinon, P., Samuels, L., and Meesawat, U. (2019) Pollen development in male sterile mangosteen (Garcinia mangostana L.) and male fertile seashore mangosteen (Garcinia celebica L.). Protoplasma 256:1545–1556.
Wang, S., Yamaguchi, M., Grienenberger, E., Martone, P.T., Samuels, A.L., Mansfield, S.D. (2019) The Class II KNOX genes KNAT3 and KNAT7 work cooperatively to influence secondary cell wall deposition and provide mechanical support to Arabidopsis stems. The Plant Journal. https://doi.org/10.1111/tpj.14541
Meents, M., Motani, S., Mansfield, S.D., Samuels, L. (2019) Organization of Xylan Production in the Golgi During Secondary Cell Wall Biosynthesis. Plant Physiology. 181: 527–546. (Featured cover article)
Livingston, S., Quilichini, T., Booth, J., Wong, D., Rensing, K., Laflamme-Yonkman, J., Castellarin, S., Bohlmann, J., Page, J., Samuels, L. (2019) Cannabis glandular trichomes alter morphology and metabolite content during flower maturation. The Plant Journal. 101: 37-56 (Featured cover article)
Perkins, M., Smith, R., Samuels, L. (2019) The Transport of Monomers during Lignification in Plants: Anything Goes but How? (invited review) Current Opinion in Plant Biotechnology, DOI: 10.1016/j.copbio.2018.09.011.
Watanabe, Y., Schneider, R., Barkwill, S., Gonzales-Vigil, E., Hill, J.L., Samuels†, A.L., Persson†, S., Mansfield†, S.D. (2018) Cellulose Synthase Complexes Display Distinct Dynamic Behaviors during Xylem Transdifferentiation. Proceedings of the National Academy of Sciences USA 115: E6366-E6374. (†=co-corresponding authors).
Chou, E., Schuetz, M., Sibout, R., Hoffman, N., Watanabe, Y., Samuels, L. (2018) Distribution, Mobility and Anchoring of Lignin-Related Oxidative Enzymes in Arabidopsis Secondary Cell Walls. Journal of Experimental Botany 69: 1849-1859.
Meents, M., Watanabe, Y., Samuels, L. (2018) The Cell Biology of Secondary Cell Wall Biosynthesis (invited review). Annals of Botany 121:1107–1125.
Smith, R.A., Schuetz, M., Karlen, S.D., Bird, D.A., Tokunaga, N., Sato, Y., Mansfield, S.D., Ralph, J., and A.L. Samuels (2017) Defining the Diverse Cell Populations Contributing to Lignification in Arabidopsis thaliana Stems. Plant Physiology 174: 1028-1036.
McFarlane H.E., Lee E.K., van Bezouwen L.S, Ross B., Rosado A., Samuels A.L. (2017) Multiscale Structural Analysis of Plant ER-PM Contact Sites. Plant and Cell Physiology 58: 478-484.
Smith, R.A., Gonzales-Vigil, E., Karlen, S., Park, J-Y., Lu, F., Wilkerson, C., Samuels, L., Ralph, J., Mansfield, S.D. (2015) Engineering Monolignol p-Coumarate Conjugates into Poplar and Arabidopsis Lignins. Plant Physiology 169: 2992-3001.
Watanabe, Y., Meents, M.J., McDonnell, L.M., Barkwill, S., Sampathkumar, A., Cartwright, H.N., Demura, T., Ehrhardt, D.W., Samuels, L. †, S. D. Mansfield† (†=co-corresponding authors). (2015) Visualization of cellulose synthases in Arabidopsis secondary cell walls. Science 350: 198-203.
Quilichini, T.D., Samuels, A.L., and Douglas, C.J. ABCG26-Mediated Polyketide Trafficking and Hydroxycinnamoyl Spermidines Contribute to Pollen Wall Exine Formation in Arabidopsis. Plant Cell doi: http://dx.doi.org/10.1105/tpc.114.130484 The Plant Cell November 2014 tpc.114.13048
Schuetz, M., Benske, A., Smith, R.E., Watanabe, Y., Tobimatsu, Y., Ralph, J., Demura, T., Ellis, B., Samuels, A.L. (2014) Laccases direct lignification in the discrete secondary cell wall domains of protoxylem. Plant Physiology. First Published on August 25, 2014; doi: http://dx.doi.org/10.1104/pp.114.245597
McFarlane, H.E., Watanabe, Y., Yang, W., Huang, Y., Ohlrogge, J., and Samuels, A.L. (2014) Golgi- and Trans-Golgi Network-Mediated Vesicle Trafficking Is Required for Wax Secretion from Epidermis. Plant Physiology 164: 1250-1260.
Quilichini T.D., Douglas C.J., and Samuels A.L. (2014) New views of tapetum ultrastructure and pollen exine development in Arabidopsis thaliana. Annals of Botany, doi: 10.1093/aob/mcu042
Smith, R.A., Schuetz, M., Roach, M., Mansfield, S.D., Ellis B.E., Samuels, L. (2013) Neighboring parenchyma cells can contribute to Arabidopsis xylem lignification, while lignification of interfascicular fibers is cell autonomous. Plant Cell 25: 3988-99.
McFarlane, H.E., Watanabe, Y., Carruthers, K., Lesvesque-Tremblay, G., Haughn, G.W., Gendre, D., Bhalerao, R.P., Samuels, A.L. (2013) The echidna mutant demonstrates that pectic polysaccharides and proteins require distinct post-Golgi vesicle traffic machinery. Plant & Cell Physiology 54: 1867-1880.
Kaneda, M. Schuetz, B.S.P. Lin, C. Chanis, B. Hamberger, T.L. Western, J. Ehlting, A.L. Samuels (2011) ABC transporters coordinately expressed during lignification of Arabidopsis stems include a set of ABCB's associated with auxin transport. Journal of Experimental Botany 62: 2063-2077.
McFarlane, H.E., Shin, J.J., Bird, D.A., and Samuels, A.L. 2010. Arabidopsis ABCG transporters, which are required for export of diverse cuticular lipids, dimerize in different combinations. Plant Cell 22: 3066-3075. [view abstract]
Quilichini, T.D., Friedmann, M.C., Samuels, A.L., and Douglas, C.J. 2010. ATP-binding cassette transporter G26 (ABCG26) is required for male fertility and pollen exine formation in Arabidopsis. Plant Physiology 154: 678-690.[view abstract]
M. Kaneda, K.H. Rensing, J.C.T. Wong, B. Banno, S.D. Mansfield, A.L. Samuels. (2008) Tracking Monolignols During Wood Development in Pinus contorta var. latifolia. Plant Physiology First published on June 11, 2008; 10.1104/pp.108.121533.
Samuels A.L., L. Kunst, R. Jetter (2008) Sealing plant surfaces: cuticular wax formation by epidermal cells. Annual Review of Plant Biology. Vol. 59: 683-70. [view full text]