Laboratory of structural membrane biochemistry
Our laboratory studies the structures of membrane proteins. Based on structure we try to understand function and what goes wrong in disease. We focus primarily on proteins in the blood-brain barrier. The long-standing question in our laboratory is how the thousands of membrane channels and transporters that exist in the cell membrane work together to help cells maintain homeostasis. With that question in mind, we study membrane proteins that are involved in nutrient, ion and water uptake, waste removal, signaling and communication.
Our laboratory is multidisciplinary. Over the last decade we have employed structural biology techniques such as electron cryo-microscopy (cryo EM), X-ray crystallography, NMR, molecular dynamics simulations, and used membrane biochemistry and biophysics to understand the function of the proteins of interest. Within electron microscopy we have published papers using electron tomography, single particle reconstructions and electron crystallography, however our specialty lies in electron diffraction.
Part of our laboratory is also devoted to method development in cryo EM. In recent years we have developed two important methods in electron diffraction, namely the fragment based phase extension and MicroED.
Some of our recent studies are outlined below.
Lysosomal amino acid transceptor SLC38A9 and mTORC pathway activation
Recent advances in intracellular amino acid transport and mechanistic target of rapamycin complex 1 (mTORC1) signaling shed light on the solute carrier 38 family A member 9 (SLC38A9), a lysosomal transporter responsible for binding and translocation of several essential amino acids. Here we present the first crystal structure of SLC38A9 from Danio rerio in complex with arginine. As captured in the cytosol-open state, the bound arginine was locked in a transitional state stabilized by the transmembrane helix 1 (TM1) of drSLC38A9 which was anchored at the grove between TM5 and 7. These anchoring interactions were mediated by the highly conserved motif WNTMM on TM1 and mutations in this motif abolished arginine transport by drSLC38A9. The underlying mechanism of substrate binding is critical for sensitizing mTORC1 signaling pathway to amino acids and for maintaining lysosomal amino acid homeostasis. This study offers a first glimpse into a prototypical model for SLC38 transporter
Using this initial structure we could compute models for representative SNAT proteins
Relevant papers for amino acid transport work
Nat. Struct. Mol. Biol., 25 (6), pp. 522–527, 2018.
The dynamic regulation of water channels
Water channels, or aquaporins, form specialized channels in membranes for water permeation. These are extremely efficient channels that allow millions of water molecules to permeate the pore per second. Because they are channels, the cell can regulate their activity dynamically to help maintain homeostasis. In the case of the eye lens water channel aquaporin-0 (AQP0), it can be regulated by at least 4 known mechanisms that we studied over the last decade. The first is irreversible and involves the cleavage of the C-terminal domain of AQP0. The cleavage results in complete pore closure and AQP0 ceases to act as a water channel. Instead it becomes an adhesive protein mediating cell-to-cell adhesive junctions (Figure 1).
Full-length AQP0 is dynamically modulated by 3 mechanisms: pH, calcium/calmodulin (Ca²⁺/CaM) and protein phosphorylation. We recently showed that the binding of Ca²⁺/CaM to AQP0 results in partial pore closure (Figure 2). The net effect is that the permeability through AQP0 halves in the presence of Ca²⁺/CaM. Conversely, we showed that phosphorylation of AQP0 by anchored PKA (AKAP2/PKA complex) abolished CaM binding, keeping AQP0 in the open conformation and functioning at maximal activity.
Our studies of channel phosphorylation led us to discover a new protein in the eye lens called AKAP2. Our biochemical and structural studies indicate that AKAPs anchor PKA onto substrate and provide the kinase a sphere of action in which the kinase could phosphorylate substrates in a cAMP independent way. This is fundamentally an exciting observation because it helps explain how fast phosphorylation can occur, as seen for example in heart cells. Moreover, we showed that inhibition of phosphorylation of AQP0 in the lens results in cataract formation. Essentially we recapitulated the lens disease ex vivo by inhibiting protein phosphorylation (Figure 3).
Membrane protein complexes
Our structure of the AQP0/CaM complex is the first for any full-length membrane channel in complex with this ubiquitous secondary messenger (Figure 4). Current efforts in the laboratory are to understand how Ca²⁺/CaM binds to and modulates the activity of other channels such as ion channels.
We are also trying to understand more about the AQP-AKAP system, in particular we are trying to assemble the AQP2-AKAP18-PKA complex and AQP0-AKAP2-PKA complex for structural studies. Intrinsically disordered regions of proteins are widespread in nature yet the mechanistic roles they play in biology are underappreciated. Such disordered segments can act simply to link functionally coupled structural domains or they can orchestrate enzymatic reactions through a variety of allosteric mechanisms. The regulatory subunits of protein kinase A provide an example of this important phenomenon where functionally defined and structurally conserved domains are connected by intrinsically disordered regions of defined length with limited sequence identity. Our studies show that this seemingly paradoxical amalgam of order and disorder permits fine-tuning of local protein phosphorylation events. The anchoring of PKA by AKAP affords the kinase a sphere of action in which multiple targets can get phosphorylated fast in a cAMP independent way (Figure 5).
Elife, 2 , pp. e01319, 2013.
Allosteric mechanism of water-channel gating by Ca²⁺-calmodulin Journal Article
Nat. Struct. Mol. Biol., 20 (9), pp. 1085–1092, 2013.
EMBO Mol Med, 4 (1), pp. 15–26, 2011.
Structure, 16 (9), pp. 1389–1398, 2008.
Nature, 438 (7068), pp. 633–638, 2005.
Aquaporin-0 Membrane Junctions Form Upon Proteolytic Cleavage Journal Article
J. Mol. Biol., 342 (4), pp. 1337–1345, 2004.
Nature, 429 (6988), pp. 193–197, 2004.
Local cAMP signaling in disease at a glance Journal Article
J. Cell. Sci., 126 (Pt 20), pp. 4537–4543, 2013.
Lipid-protein interactions probed by electron crystallography Journal Article
Curr. Opin. Struct. Biol., 19 (5), pp. 560–565, 2009.
Electron Crystallography of Aquaporins Journal Article
IUBMB Life, 60 (7), pp. 430–436, 2008.
Junction-forming aquaporins Journal Article
Curr. Opin. Struct. Biol., 18 (2), pp. 229–235, 2008.
The structure of aquaporins Journal Article
Q. Rev. Biophys., 39 (4), pp. 361–396, 2006.
Membrane transporters involved in nutrient uptake
The major facilitator superfamily of membrane proteins is the largest collection of structurally related membrane proteins that transport a wide array of substrates. The proton-coupled sugar transporter XylE is the first member of the MFS that has been captured and structurally characterized in multiple transporting conformations including both the outward and inward facing states. We determined the crystal structure of XylE in a new inward-facing open conformation. Structural comparison of XylE in this conformation with its outward-facing partially occluded conformation reveals how this transporter functions through a non-symmetrical rocker switch movement of the N-domain as a rigid body and the C-domain as a flexible body. Molecular dynamics simulations were employed to help describe how XylE transitions in a lipid membrane to facilitate sugar transport. (Figure 6)
Nitrate is the preferred nitrogen source for plants on which all higher forms of life ultimately depend. Plants and microorganisms evolved to efficiently assimilate nitrate. Despite decades of effort no structure was available for any nitrate transport protein and the mechanism by which nitrate is transported remained largely obscure until our study was published. We reported the structure of a bacterial nitrate/nitrite transport protein, NarK, from Escherichia coli, with and without substrate. The structures revealed a positively charged substrate-translocation pathway lacking protonatable residues, suggesting that NarK functions as a nitrate/nitrite exchanger and that H⁺s are unlikely to be co-transported. Conserved arginine residues form the substrate-binding pocket, which is formed by association of helices from the two halves of NarK. Key residues that are important for substrate recognition and transport were identified and related to extensive mutagenesis and functional studies. We proposed that NarK exchanges nitrate for nitrite by a rocker-switch mechanism facilitated by inter-domain H-bond networks. (Figure 7)
Amino acid uptake
The amino acid, polyamine, and organocation (APC) superfamily is the second largest superfamily of membrane proteins forming secondary transporters that move a range of organic molecules across the cell membrane and that can transport both D- and L- amino acids. Here we report two crystal structures of an APC member from Methanococcus maripaludis, the alanine or glycine:cation symporter (AgcS), with L- and D- alanine. Structural analysis combined with site-directed mutagenesis and functional studies inform on substrate binding, specificity, and modulation of the AgcS family and reveal key structural features that allow this transporter to accommodate glycine and both L- and D- type alanine while excluding all other amino acids. Mutation of key residues in the substrate binding site expand the transporters selectivity to include valine and leucine. Moreover, as a transporter that binds both enantiomers of alanine, the present structures provide an unprecedented opportunity to gain insights into the mechanism of stereo-selectivity in membrane transporters.
Nat Commun, 5 , pp. 4521, 2014.
Crystal structure of a nitrate/nitrite exchanger Journal Article
Nature, 497 (7451), pp. 647–651, 2013.
Structure, 18 (11), pp. 1512–1521, 2010.
J. Mol. Biol., 396 (3), pp. 593–601, 2010.
Method development in cryo‐EM
Fragment‐based phase extension
In electron crystallography membrane protein structure is determined from two-dimensional crystals where the protein is embedded in a membrane. Once large and well-ordered 2D crystals are grown one of the bottlenecks in electron crystallography is the collection of image data to directly provide experimental phases to high resolution. We developed a new approach to bypass this bottleneck, eliminating the need for high-resolution imaging. We used the strengths of electron crystallography in rapidly obtaining accurate experimental phase information from low-resolution images and accurate high-resolution amplitude information from electron diffraction. The low-resolution experimental phases were used for the placement of α-helix fragments and extended to high resolution using phases from the fragments. Phases were further improved by density modifications followed by fragment expansion and structure refinement against the high-resolution diffraction data. Using this approach, structures of three membrane proteins were determined rapidly and accurately to atomic resolution without high-resolution image data. (Figure 8)
MicroED —Three‐dimensional electron crystallography of protein microcrystals
We demonstrated that it is feasible to determine high-resolution protein structures by electron crystallography of three-dimensional crystals in an electron cryo-microscope (CryoEM). Lysozyme microcrystals were frozen on an electron microscopy grid, and electron diffraction data collected to 1.7Å resolution. We developed a data collection protocol to collect a full-tilt series in electron diffraction to atomic resolution. A single tilt series contains up to 90 individual diffraction patterns collected from a single crystal with tilt angle increment of 0.1 – 1° and a total accumulated electron dose less than 10 electrons per angstrom squared. We indexed the data from three crystals and used them for structure determination of lysozyme by molecular replacement followed by crystallographic refinement to 2.9Å resolution (Figure 9). This proof of principle paves the way for the implementation of a new technique, which we name “MicroED”, that may have wide applicability in structural biology. Current efforts include new phasing methods, automation and program development.
An example of lysozyme MicroED data can be viewed here.
In 2014 we further inmproved the MicroED method. Firstly, we developed an improved data collection protocol for MicroED called Continuous rotation. Microcrystals are continuously rotated during data collection yielding improved data, and allowing data processing with the crystallographic software tool MOSFLM, resulting in improved resolution for the model protein lysozyme to 2.5Å resolution. These improvements pave the way for the broad implementation and application of MicroED in structural biology. Current efforts include new phasing methods, automation and program development.
Secondly, we used the improved MicroED protocols for data collection and analysis to determine the structure of catalase. Bovine liver catalase crystals that were only ~160nm thick were used for the structure analysis. A single crystal yielded data to 3.2Å resolution enabling structure determination rapidly.
An example of catalase MicroED data can be viewed here.
In 2015 we published the first two previously unknown structures determined by MicroED. The structures of two peptides from the toxic core of a-synuclein of Parkinsons’ Disease. The structures were determined from vanishingly small crystals, only ~200nm thick and wide, and yielded 1.4Å resolution. These structures, which are currently the highest resolution structures determined to date by any cryo EM method, show new and important structural information that could aid in the development of pharmaceuticals against this devastating neurological disease. The study, which was published by Nature also show a number of protons for the very first time.
Follow MicroED on Twitter #MicroED
J Appl Crystallogr, 49 (Pt 3), pp. 1029–1034, 2016.
Nat Protoc, 11 (5), pp. 895–904, 2016.
Nature, 525 (7570), pp. 486–490, 2015.
MicroED data collection and processing Journal Article
Acta Crystallogr A Found Adv, 71 (Pt 4), pp. 353–360, 2015.
Structure of catalase determined by MicroED Journal Article
Elife, 3 , pp. e03600, 2014.
Nat. Methods, 11 (9), pp. 927–930, 2014.
J Appl Crystallogr, 47 (Pt 3), pp. 1140–1145, 2014.
Elife, 2 , pp. e01345, 2013.
Structure, 19 (7), pp. 976–987, 2011.
Relevant Reviews and Book Chapters:
Protein structure determination by MicroED Journal Article
Curr. Opin. Struct. Biol., 27 , pp. 24–31, 2014.
Curr Protoc Protein Sci, 72 (1), pp. 17.15.1–17.15.11, 2013.
Electron Crystallography of Soluble and Membrane Proteins, 955 , Chapter 14, pp. 243–272, 2012.
Electron Crystallography of Soluble and Membrane Proteins, 955 , Chapter 9, pp. 153–169, 2012.
High-Throughput Methods for Electron Crystallography Book Chapter
Electron Crystallography of Soluble and Membrane Proteins, 955 , Chapter 15, pp. 273–296, 2012.
Structure, 19 (10), pp. 1381–1393, 2011.
Computational design of genetically encoded self‐assembling proteins
In collaboration with David Baker (HHMI, UW) we are designing genetically encoded self assembling proteins for cellular microcircuitry.
We describe a general computational method for designing proteins that self-assemble to a desired symmetric architecture. Protein building blocks are docked together symmetrically to identify complementary packing arrangements, and low-energy protein-protein interfaces are then designed between the building blocks in order to drive self-assembly. Here we use trimeric protein building blocks to design a 24-subunit, 13 nm diameter complex with octahedral symmetry and two related variants of a 12-subunit, 11 nm diameter complex with tetrahedral symmetry. The designed proteins assembled to the desired oligomeric states in solution, and crystal structures of the complexes revealed that the resulting materials closely match the design models. The method can be used to design a wide variety of self-assembling protein nanomaterials. (Figure 10)
Protein Sci., 24 (10), pp. 1695–1701, 2015.
Science, 348 (6241), pp. 1365–1368, 2015.
Nature, 510 (7503), pp. 103–108, 2014.
Science, 336 (6085), pp. 1171–1174, 2012.
Electrophysiology—channel recordings toward structure‐function analysis
We use electrophysiology and patch clamping techniques to study the function of channels and transporters. We use the Xenopus oocyte expression system as well as whole cell patch but we also plan to record channel function from highly ordered two-dimensional crystals for a direct correlation between structure and function of target proteins as they are embedded within a biological membrane.
Other notable studies (not currently active in the lab)
Structure of the vibrio cholera toxin secretion channel
In collaboration with Wim Hol (UW) we studied the structure of the vibrio cholera toxin secretion channel.
The type II secretion system (T2SS) is a macromolecular complex spanning the bacterial inner and outer membranes of Gram-negative bacteria, including many pathogenic bacteria such as Vibrio cholerae and enterotoxigenic Escherichia coli. The T2SS secretes folded proteins including cholera toxin and heat-labile enterotoxin. The major outer membrane T2SS protein is the “secretin” GspD. Electron cryomicroscopy (cryoEM) reconstruction of the V. cholerae secretin at 19 Å resolution reveals a dodecameric structure reminiscent of a barrel with a large channel at its center that appears to be in a closed state. On the periplasmic side of the channel vestibule contains both a constriction and a gate. On the extracellular side a large chamber is enclosed by a cap structure. By combining our results with structural data on a large exoprotein and the dimensions of the tip of the pseudopilus of the T2SS, we provide a structural basis for a possible secretion mechanism of exoproteins by the T2SS in which the constriction site plays a critical role. (Figure 12)
Channels (Austin), 5 (3), pp. 215–218, 2011.
Nat. Struct. Mol. Biol., 17 (10), pp. 1226–1232, 2010.
Trends Biochem. Sci., 36 (8), pp. 433–443, 2011.
Structure and function of the yeast kinetochore and microtubule dynamics
In collaboration with Sue Biggins (FHCRC) we studied the structure of the yeast kinetochore by electron tomography. In collaboration with Trisha Davis and Chip Asbury (UW) we studied microtubule dynamics and microtubule binding proteins.
Chromosomes must be accurately partitioned to daughter cells to prevent genomic instability and aneuploidy, a hallmark of many tumors and birth defects. Kinetochores are macromolecular machines that move chromosomes by maintaining load-bearing attachments to the assembling and disassembling tips of spindle microtubules. The mechanism by which kinetochores attach to microtubules is still not clear although a number of models have been proposed. Sues laboratory previously developed an assay to purify functional native budding yeast kinetochore particles that contain the majority of core structural components and can maintain attachments to microtubules under force. We presented the structure of these isolated kinetochore particles as visualized by electron microscopy (EM) and electron tomography of negatively stained preparations. The budding yeast kinetochore appeared as a ~126 nm particle having a large central hub attached to multiple outer globular domains. Microtubule binding experiments indicated that the globular domains are important for microtubule attachments both in the presence or absence of a ring encircling the microtubule. Our data showed that kinetochores bind to microtubules via multivalent attachments, consistent with a biased diffusion mechanism where multiple kinetochore components cooperate to form a strong yet dynamic linkage to the microtubule. Although rings are not required for lateral binding, they likely maintain processive attachments to the ends of dynamic microtubules. These studies lay the foundation to uncover the key mechanical and regulatory mechanisms by which kinetochores control chromosome segregation and cell division. (Figure 13)
Proc. Natl. Acad. Sci. U.S.A., 109 (40), pp. 16113–16118, 2012.
Nat. Struct. Mol. Biol., 19 (9), pp. 925–929, 2012.
Nature, 468 (7323), pp. 576–579, 2010.
J. Cell Biol., 189 (4), pp. 713–723, 2010.
Nat. Cell Biol., 9 (7), pp. 832–837, 2007.
Updated June 22 2018 © Tamir Gonen