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Our Research

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.

 

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).

 

 

Relevant papers:

 

1. Gonen T., Sliz P., Cheng Y., Kistler J. and Walz T. (2004) Aquaporin-0 membrane junctions reveal the

structure of a closed water pore. Nature. 429 : 193 – 197.

2. Gonen T., Cheng Y., Kistler J. and Walz T. (2004) Aquaporin-0 membrane junctions form upon

proteolytic cleavage. Journal of Molecular Biology 342 : 1337 - 1345.

3. Gonen T., Cheng Y., Sliz P., Hiroaki Y., Fujiyoshi Y., Harrison SC., Walz T (2005) Lipid–protein

interactions in double layered two dimensional crystals of AQP0. Nature 438: 633 - 638.

4. Reichow L.S. and Gonen T*. (2008) Non-canonical binding of calmodulin to aquaporin-0:

implications for channel regulation. Structure. 16: 1389 – 1398.

5. Gold MG, Reichow SL., O’Neill SE, Weisbrod CR, Langeberg LK, Bruce JE, Gonen T* and Scott

JD*. (2012) AKAP2 anchors PKA with aquaporin-0 to support ocular lens transparency.

EMBO Molecular Medicine 4: 15-26.

6. Reichow S.L, Clemens D.M., Freites J.A., Németh-Cahalan K.L., Heyden M., Tobias D.J., Hall J.E*.

and Gonen T*. (2013) Allosteric mechanism of water channel gating by Ca2+–calmodulin. Nature

Structural and Molecular Biology – 20 (9): 1085 - 1092.

7. Smith FD., Reichow LS., Esseltine JL., Shi D., Langeberg LK., Scott J* and Gonen T* (2013). Intrinsic

disorder within an AKAP-Protein kinase A complex guides local substrate phosphorylation. eLife 2:e01319.

 

Relevant Reviews:

1. Gonen T*. and Walz T* (2006) The structure of Aquaporins. Quarterly Reviews in Biophysics 39 :

361 -396.

2. Engel, A., Fujiyoshi, Y., Gonen, T. and Walz, T. (2008) Junction forming aquaporins. Current

Opinion in Structural Biology. 18 : 229 - 235.

3. Andrews, S.A., Reichow, L.S. and Gonen T*. (2008) Electron crystallography of aquaporins.

IUBMB Life 60: 430 – 436.

4. Reichow, S.L. and Gonen T*. (2009) Lipid-protein interactions probed by electron crystallography.

Current Opinion in Structural Biology. 19: 560 - 565

5. Gold M. Gonen T and Scott J. (2013) Local cAMP signaling in disease at a glance. Journal of Cell

Science. 126: 4537 - 4543.

 

 

 

Membrane transporters involved in nutrient uptake

 

Sugar 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)

 

 

Nitrogen uptake

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)

 

 

Relevant papers

1. Zheng H., Taraska J., Merz A. and Gonen T*. (2010) The prototypical H+/galactose symporter GalP

assembles into functional trimers. Journal of Molecular Biology. 396 : 593 – 601.

2. Wisedchaisri G., Dranow D.M., Lie T.J., Bonanno J.B., Patskovsky Y., Ozyurt S.A., Michael Sauder

J.M., Almo S.C., Wasserman S.R., Burley S.K., Leigh J.A. and Gonen T*. (2010) Structural

underpinnings of nitrogen regulation by the prototypical nitrogen-responsive transcriptional factor

NrpR. Structure 18 1512 - 1521.

3. Zheng H, Wisedchaisri W and Gonen T*. (2013) Crystal structure of a nitrate/nitrite exchanger.

Nature – 497: 647-651.

4. Wisedchaisri G., Park MS., Iadanza MG., Zheng H. and Gonen T*. (2014) Proton-coupled sugar transport in

               the prototypical major facilitator superfamily protein XylE. Nature Communications – (5)4521: 1 - 11.

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

 

 

Relevant papers

1. Wisedchaisri G and Gonen T*. (2011) Fragment based phase extension for membrane protein

structure determination by electron crystallography. Structure 19: 976 - 987.

2. Shi D., Nannenga B., Iadanza MG. and Gonen T* (2013). MicroED – Three dimensional electron

crystallography of protein microcrystals. eLife 2:e01345: 1 - 17.

3. Iadanza MG. and Gonen T*. A suite of software for processing MicroED data of extremely small protein

crystals. Journal of Applied Crystallography. 47: 1140 – 1145.

4.Nannenga BL, Shi D., Leslie AGW. and Gonen T* (2014). High-resolution structure determination by

continuous rotation data collection in MicroED. Nature Methods 11 (9): 927 – 930.

5. Nannenga BL, Shi D., Hattne J., Reyes F. and Gonen T* (2014). Structure of catalase determined by

MicroED. eLife 3:e03600: 1 – 11.

6. Hattne J., Reyes FE., Nannenga BL., Shi D., de la Cruz J., Leslie AGW. And Gonen T*. MicroED data

collection and processing. Acta Crystallographica section A. A71: 353 - 360 (2015).

7. Rodriguez A.J., Ivanova M., Sawaya MR., Cascio D., Reyes F., Shi D., Sangwan S., Guenther EL.,

Johnson L, Zhang M., Jiang L., Arbing M., Nannega B., Hattne J., Whitelegge J., Brewster AS.,

Messerschmidt M., Boutet S., Sauter NK., Gonen T* and Eisenberg D* Structure of the toxic core

of a-synuclein from invisible crystals. Nature 525 (7570): 486 - 490 (2015).

8. Hattne J., Shi D., de la Cruz J., Reyes FE., and Gonen T*. Modeling truncated intensities of faint

reflections in MicroED images. Journal of Applied Crystallography - In Press (2016).

9. Shi D., Nannenga B., de la Cruz J., Jiu L., Guillermo C., Hattne J., Reyes FE., Sawtelle S. and

Gonen T*. The collection of MicroED data for macromolecular crystallography. Nature Protocols 11 (5) : 895 - 904 (2016).

 

Relevant Reviews and Book Chapters:

1. Wisedchaisri W., Reichow S.L. and Gonen T*. (2011) Advances in structural and functional analysis

of membrane proteins by electron crystallography. Structure. 19:1381-93.

2. Wisedchaisri W. and Gonen T*. (2013) Phasing Electron Diffraction Data by Molecular Replacement:

Strategy for Structure Determination and Refinement. Methods in Molecular Biology 955: 243 – 272.

3. Gonen T*. (2013) The collection of high-resolution electron diffraction data. Methods in Molecular

Biology 955: 153 – 169.

4. Stokes D, Ubarretxena I, Gonen T and Engel A. (2013) High throughout methods in electron

crystallography. Methods in Molecular Biology 955: 273 – 296.

5. Nannenga B, Iadanza M, Vollmar B and Gonen T*. (2013) Electron crystallography of membrane

proteins: crystallization and screening strategies using negative stain electron microscopy.

Current Protocols in Protein Science – 17 (15): 1-11.

6. Nannenga BL. and Gonen T*. Protein structure determination by MicroED. Current Opinion in

Structural Biology. 27: 24 - 31 (2014).

 

 


 

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)

 

Relevant papers

1. King NP., Sheffler W., Sawaya MR., Vollmar BS., Sumida JP., Andre I., Gonen T., Yeates TO. And Baker D. (2012) Computational design of self-assembling protein nanomaterials with atomic level accuracy. Science. 336: 1171 – 1174.

2. King NP, Bale J, Sheffler W, McNamara DE., Gonen S., Gonen T., Yeates TO. and Baker D. Accurate design of coassembling multi-component protein nanomaterials. Nature 510 (7503): 103 – 108.

3. Bale JB., Park RU., Liu Y., Gonen S., Gonen T., Cascio D., King NP., Yeates TO., and Baker D (2015) Structure of a designed tetrahedral protein assembly variant engineered to have improved soluble expression. Protein Science – In Press.

4. Gonen S., DiMIao F., Gonen T* and Baker D*. Design of ordered two-dimensional arrays mediated by noncovalent protein-protein interfaces. Science 348 (6241): 1365 - 1368 (2015)

 


 

 

 

Electrophysiology: channel recordings toward structure-function nalysis

 

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.

(Figure 11)

 

 

 

 

 

 

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)

 

Relevant papers:

1. Reichow S.L., Korotkov K.V., Hol W.G.J*. and Gonen T*. (2010) Structure of the cholera toxin

secretion channel in its closed state. Nature Structural & Molecular Biology. 17 : 1226 - 1232.

2. Reichow SL., Korotkov KV., Gonen M., Sun J., Delarosa J., Hol WGJ* and Gonen T* (2011) The

binding of cholera toxin to the periplasmic vestibule of the type II secretion channel. Channels 5 (3): 215 – 218.

 

Relevant Reviews:

1. Korotkov KV., Gonen T and Hol WGJ. Secretins: dynamic channels for protein transport across

membranes. Trends in Biochemical Sciences 36 (8) : 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)

 

 

Relevant papers

1. Franck, A.D., Powers, A.F., Gestaut, D.R., Gonen, T., Davis, T.N. and Asbury, C.L. (2007) Tension

applied through the Dam1 complex promotes microtubule elongation providing a direct mechanism for length control in mitosis. Nature Cell Biology. 9 : 832 - 837.

2. Tien J.F., Umbreit N.T., Gestaut D.R., Franck A.D., Cooper J., Wordeman L., Gonen T., Asbury C.L.

and Davis T.N. (2010) Cooperation of the Dam1 and Ndc80 kinetochore complexes enhances processive microtubule coupling and is regulated by Aurora B. Journal of Cell Biology. 189 : 713 – 723.

3. Akiyoshi B., Sarangapani K.K., Powers A.F., Nelson C.R., Reichow S.L., Arellano-Santoyo H.,

Gonen T., Ranish J.A., Asbury C.L., and Biggins S. (2010) Tension directly stabilizes reconstituted kinetochore-microtubule attachments. Nature 468 : 576 - 581.

4. Umbreita NT., Gestaut DR., Tien JF., Vollmar BS., Gonen T, Asbury CL and Davis TN. (2012) The

Ndc80 kinetochore complex directly modulates microtubule dynamics. Proceedings of the National Academy of Sciences 109 (40): 16113 - 16118.

5. Gonen S., Akiyoshi B., Iadanza MG., Shi D., Duggan N., Biggins S*. and Gonen T*. (2012) The

structure of purified kinetochores reveals multiple microtubule-attachment sites. Nature Structural and Molecular Biology19 (9): 925 – 930.

 

Updated May 15 2016 © Tamir Gonen