Principal Investigator: Rajendra K. Agrawal, Ph.D.

Protein synthesis (or translation) is a fundamental process in all forms of life. An understanding of the molecular mechanisms underlying the various steps of translation is of seminal importance in a large range of fields in Biology.

We study the structure and function of ribosome, the protein synthesizing machinery of the cell and cellular organelles such as mitochondria, using cryo-EM single-particle reconstruction techniques (Frank et al., 2000; Shaikh at al., 2008). The current focus in the Agrawal laboratory is to structurally characterize the mammalian mitochondrial translation by studying molecular interactions between the mitochondrial ribosome and its ligands during all steps of protein synthesis, at 5-8 Å resolution in each case. The goal is to come to an understanding of (i) the unique features of the organellar ribosmes in general, and mitochondrial ribosomes in particular, as molecular machines, and (ii) their structural evolution from the primitive endosymbiotic bacteria, over the course of 2 billion years, to its current form within the mitochondrion of various cell types/organisms.

Cryo-EM single-particle reconstruction has evolved to become the most powerful approach to study ligand binding and conformational changes accompanying translation. Recent reviews (Agrawal et al., 2011, Agrawal and Sharma, 2012; Frank, 2012) gives only a glimpse of many results that have accumulated since 1996, when tRNA binding to the ribosome was first visualized (Agrawal et al., 1996). The EF-G mediated tRNA translocation was also first visualized by cryo-EM (Agrawal et al., 1998; 1999; Frank and Agrawal, 2000), and then main sequence of events during the translation elongation cycle were visualized (Agrawal et al., 2000). These early studies laid much of the groundwork for several functional ligand-binding studies that subsequently followed during the past decade in the Agrawal laboratory and many other laboratories. The Agrawal laboratory obtained the first cryo-EM structures of (i) the mammalian mitochondrial ribosome (Sharma et al., 2003), (ii) chloroplast ribosome (Sharma et al., 2007), and (iii) a protistan mitochondrial ribosome (Sharma et al., 2009). These studies revealed several unique features of the organellar translational machinery as reviewed recently (Agrawal and Sharma, 2012; Agrawal et al., 2011). In addition, the Agrawal laboratory continues to study specific steps of cytoplasmic translation, for example, RRF- and EF-G-dependent ribosome recycling (Agrawal et al., 2004; Barat et al., 2007; Yokoyama et al., 2012), in order to understand the underlying molecular mechanisms of the process and to compare the analogous mechanisms in the mitochondrial system. Using existing tools of cryo-EM single-particle reconstruction, and tools being developed for time-resolved cryo-EM studies (e.g., Lu et al., 2009), we are pursuing studies to unravel detailed molecular mechanism of various steps of mammalian mitochondrial translation at 5-8 Å resolution. Here are examples from some of the more recently published works:

Figure 1. Comparison between the segmented cryo-EM maps of ribosomes from (A) bacteria (70S, E. coli), (B) chloroplasts (70S, Spinacea oleracea), (C) protistan mitochondria (50S, Leishmania tarentolae), and (D) mammalian mitochondria (55S, Bos taurus). Ribosomal Proteins of small and large ribosomal subunits are colored yellow and blue, respectively, while ribosomal RNAs are colored orange and magenta, respectively. Results revealed that mitochondrial ribosome surfaces are shielded extensively by protein masses. (E, F) Secondary structures of small and large ribosomal subunit RNAs, respectively, from bacteria (grey), mammalian mitochondria (green), and L. tarentolae mitochondria (red). Structures of mitochondrial rRNAs were derived from analyses of respective cryo-EM maps.
 

Figure 2. A side-by-side comparison of topographies of the nascent polypeptide-exit sites in 3D cryo-electron microscopic maps of the ribosome from (A) bacteria (E. coli), (B) chloroplasts (S. oleracea), (C) protozoan (L. tarentolae) mitochondria, and (D) mammalian (B. taurus) mitochondria, suggesting that the mechanism of nascent polypeptide (red) exit in mitochondria is dramatically different from those in bacteria and chloroplasts. Pink and green are conserved rRNAs and proteins, respectively. Solid blue masses represent organelle specific protein masses.
 

Figure 3. Structure of the mammalian mitochondrial translation initiation factor 2 (IF2mt) as derived by single-particle cryo-EM, and location of its mitochondrial-specific insertion domain relative to the binding site of bacterial IF1. (A) Domain alignment of E. coli IF2 and bovine IF2mt. The mature IF2mt (i.e., after deletion of the mitochondrial import sequence) starts from aa residue 78. The 37-aa insertion domain in IF2mt is highlighted in red. (B) The cryo-EM map of the E. coli 70S ribosome (30S subunit, yellow; 50S subunit, blue) in complex with IF2mt (red), initiator tRNA (green) at the P/I position (P/I stands for P-site tRNA at the initiator position). (C) Fitting of atomic models of the IF2mt and initiator tRNA into the corresponding cryo-EM densities (meshwork) extracted from the map shown in panel (B). The color codes used for various domains of IF2mt are the same as in panel A. Asterisk (*) point to the region that would correspond to domain III of IF2mt but was not modeled. (D) Binding positions of the insertion domain (red), and (E) IF1 (green), into a common binding pocket on the SSU of the ribosome. Components of the binding pocket on SSU, the 16S ribosomal RNA helices (h18 and h44) and ribosomal protein S12, are identified.

Figure 4. Segmented cryo-EM maps of the bacterial 70S ribosome post-termination complex (PoTC) in complex with ttRRF and in complex with ttRRF and EF-G. (A) The PoTC•ttRRF complex. The 70S ribosome (yellow, 30S subunit; and blue, 50S subunit), with ttRRF (red); (B) The PoTC•ttRRF•EF-G•GDP•FA complex, shown in a matching view as in panel A. Densities for both ttRRF (red) and ecEF-G (orange) are readily identifiable. Position occupied by domain II of ttRRF in panels A is now occupied by domain V of EF-G. In both complexes density corresponding to mRNA (dark blue) can also be seen. (C) Molecular interactions between the ribosome and ttRRF in the PoTC•ttRRF complex. (D) Molecular interactions between ttRRF and EF-G within the PoTC•ttRRF•EF-G•GDP•FA complex. Thumbnails to the lower left of panel C and lower right of panel D depict overall orientations of the ribosome in those panels.

Principal Investigator: Haixin Sui, Ph.D.

Microtubules and microtubule-based complexes are essential in all types of cells. Molecular motors on microtubules are responsible for movement of cells and within cells, from mitosis to muscle action, as well for transport of materials within cells, and they form part of the cell’s cytoskeletal scaffold. Members of our group are studying three aspects of microtubules: (1) structure and function of microtubule-based motors, (2) assembly, maintenance, and regulation of microtubule-based complexes, and (3) interaction of microtubules with the kinetochore during mitosis.

Microtubule complexes: The Haixin Sui laboratory studies functional mechanisms of cytoskeleton assemblies in cells and organelles using structural imaging methods. Through cryo-EM and image processing, the lab has achieved high-quality structural maps for microtubule singlets of multiple structural types. We have also obtained the molecular architecture of microtubule doublets. These results have provided structural insight into their assembly and mechanical properties.

Microtubules (singlets): The microtubules are critically important in maintaining shapes of cells and organelles (cilia), and in ensuring correct dell division. They also serve as the railways for motor-based intracellular transportation. All these functions induce mechanical stress on microtubule structure and require a combination of rigidity and flexibility. The high-quality maps of various types of microtubules have been achieved at a resolution better than 9A, which enabled us to construct reliable structural models (Figure 1). Comparison of these structures offers the following insight into the lateral deformation and bending property of microtubules. (1) The loop-loop interaction offers flexibility for microtubule lateral deformation. (2) The molecular configuration of protofilament lateral interaction limits the degree of lateral deformation. (3) Local structures surrounding the inter-protofilament interaction stabilize the loop interactions and contribute to the rigidity of microtubules. (4) The seam is not significant in the mechanical properties of microtubules (Sui and Downing, 2010).

Figure 1. Structures of various types of microtubules: (a) Density maps of different structural types of microtubules containing protofilament numbers of 11 (pink), 12 (orange), 13 (yellow), 14 (green), 15 (blue), 16 (purple). These density maps were reconstructed from cryo-electron micrographs of microtubule mixtures obtained by polymerization of purified tubulin. The background image represents the typical morphology of the frozen-hydrated microtubule mixture with microtubules of various diameters. (b) The density maps clearly revealed secondary structural features and enabled constructing structural models of microtubules with good accuracy.

Microtubule doublets: Microtubule doublets are usually found in arrays of nine doublets and serve as the major framework in eukaryotic cilia or flagella. This framework contributes to the elastic property required for the mechano-sensory function of primary cilia and the beating function of motile cilia. Both microtubule singlets and doublets are mainly composed of tubulin, but the doublets also incorporate a number of other structural-protein components. Using cryo-electron tomography and image averaging, we obtained a 3D structural map of intact microtubule doublets at a resolution better than 3 nm (Figure 2). The map disclosed an 8% lateral deformation of the complete microtubule (A-tubule) in the doublet. The deformation suggests a flexibility of inter-protofilament interaction, which allows the microtubules to undergo a certain degree of lateral deformation. The map also revealed protein densities bound to the inside of the microtubule. These proteins, named intra-luminal microtubule-associated proteins (iMAPs; Amos 2010), are believed to play important roles in stabilizing or modifying inter-protofilament interactions and contribute to the lateral deformation of microtubules (Sui and Downing, 2006; Downing and Sui 2007).

Figure 2. Microtubule doublet: In the left panel, the receding gray-scale image is one of the images in one tomographic tilt series. The image just below is a vertical section through the 3-D reconstruction generated from this series. The density in gold color is the resultant 3D map of microtubule doublets with the tubulin structural model fitted into it. In the middle and right panels, the density map reveals an 8% lateral distortion of the complete microtubule (A-tubule). Protein densities other than tubulins, colored yellow, are clearly revealed inside the microtubule doublets. Inside the A-tubule, intraluminal microtubule- associated proteins (iMAPs) are visualized on the intraluminal wall, where the inter-protofilament interactions locate for integration of the microtubule structure.

Primary cilium assembly and maintenance

Primary cilia are non-motile antenna-like sensors that protrude into extracellular space for detecting a wide range of signals, including signals for cell proliferation control. Problems in primary cilium assembly and structure/length maintenance, and failures to distribute ciliary sensory proteins to specific locations of the cilium, are linked to a wide variety of medical disorders and developmental abnormalities (termed ciliopathies). Assembly and maintenance of the ciliary framework (axoneme) and distribution of sensory proteins in the primary cilium rely on motor-driven intraflagellar transport (IFT) trains that travel along axonemal microtubule complexes (MtCs). Knowing details about 3D structure of primary cilium and the IFT processes in the primary cilium is an essential foundation for understanding the structural regulation of the primary cilia and the proper distribution of sensory proteins in the cilia.

We have obtained 3D structure of primary cilia of kidney cells by serial sectional electron tomography (Figure 3). Surprisingly, we found the primary cilium structure deviated greatly from the commonly acknowledged 9+0 paradigm. The 3D structure of primary cilia raised some new questions on the fundamental understandings of IFT process. In our research, we are applying an innovative strategy that combines 3D cell culturing methods and light microscopy with techniques of 3D-EM specimen preparation and data analysis to overcome experimental difficulties for studying primary cilia of epithelial cells and their IFTs.

Figure 3. Overview of the primary cilium structure: The typical primary cilium structural map obtained by combining 33 serial dual-axis tomograms. (a) The structural model built from 3D structural map obtained by serial tomograms. Progressive changes of the microtubule configuration from proximal to distal end of the cilium are demonstrated by the end-on cross-section views shown in (b) – (i) that correspond to the locations marked in (a). The viewing direction is from the basal to the distal end. The color key is shown at the bottom.

Principal Investigator: Haixin Sui, Ph.D.

Correlative light and electron microscopy (CLEM) combines dynamic information from fluorescence light microscopy (LM) with ultrastructural information from electron microscopy (EM) of the same biological specimen. It is a powerful approach to addressing fundamental questions that neither LM nor EM alone is able to answer, revealing functional mechanisms at molecular and structural levels to advance disease mechanism understanding and guide new drug and therapeutic treatment development. Our goal is to develop convenient tools and methods to simplify CLEM studies for cells, tissues or mini-organs (i.e., organoids). In collaboration with Prof. Xie, we utilize techniques in microfabrication to make designated substrates for convenient CLEM studies of a broad range of cells and thin tissues in biomedical research.

Principal Investigator: Rajendra K. Agrawal, Ph.D.

The goal of structural biology is to understand physiological function from three-dimensional organization. Electron microscopy (EM) is one general technique to obtain a 3D structure. Whereas the end product of conventional EM is a static structure, it would be informative to observe a structure in a sequence of states in order to understand how it works. Our goal is to develop methods to initiate reactions involving macromolecules, trap them in intermediate states, and visualize them using electron microscopy. We have implemented three modes to prepare time-resolved samples for EM but current development is concentrated on the first mode: a Microfluidic Mixer-Sprayer

Microfluidic Mixer-Sprayer: We have developed a novel integrated microfluidic device that allows small volumes of two reactants to be mixed in less than 1 ms, react for a time set by the device, and then sprayed onto a dry EM grid as it is being plunged into cryogen (Lu et al. 2009, 2010; Barnard et al. 2009). Several devices have been constructed that allow reaction times ranging from 10 – 500 ms to be investigated. The current configurations require about 10 microliters of each macromolecular reactant per grid at a concentration in the micromolar range.

Figure 1. Micromixer development. (a) Configuration of the micromixer, with detailed dimensions labeled. SEM images on the bottom show the mixing channel as fabricated using DRIE etching technology (left), and the details of half of the butterfly mixing unit (right). (b) Fluorescence microscopy images of mixing fluorescein and Fluoro Sphere Red in the fabricated mixer at various flow rates. (c) Experimental and simulated mixing indices showing essentially complete mixing at flow rates above 4 μL/sec. (d) Activity of malate dehydrogenase is preserved at various flow rates.

Figure 2. Experimental setup with microspray generation shown: (a) Diagram of the experimental system, including syringe pump for liquid transport, monolithic device for mixing and spraying, and cryo-EM grid for droplet collection. (b) Photo of the entire experimental system setup. (c) Close-up of the device with fittings, which is mounted on a plastic holder. A tweezers (connected to the plunger) with an EM grid placed at the exit of the spray nozzle. (d) Fine microspray (blue arrows) generated by the device with the liquid flow rate at 6.0 μL/s, and with nitrogen gas pressure at 50 psi (set by nitrogen tank regulator).

Figure 3. Monolithic microfluidic devices for mixing and spraying (a) Detailed schematic of monolithic device (micromixer/sprayer) design showing (1) device design layout, (2) micromixer configuration, (3) SEM image of the mixer, (4) external atomization nozzle design, (5) SEM image of the nozzle, (6) device top view, and (7) device bottom view. (b) The droplet size distributions for two of the devices with the external atomization nozzles, showing the droplet size mainly in the range of 10-30 μm.

Figure 4. Cryo-EM of ribosomes and ferritin that were mixed and sprayed by the monolithic device #2. Upper left: low magnification micrograph showing the edge of a droplet that has spread sufficiently thin for image. Right: High magnification view of a hole containing thin ice and showing presence of both ferritin (dark structures) and ribosomes (lighter structures). Lower left: Surface representation of a three-dimensional reconstruction of the ribosome determined from images such as that shown in B. Resolution is 18.9Å. [Adapted from Lu et al. J. Struct. Biol., 2009]

Figure 5. Classification of ~4400 partially re-associated 70S images (out of 17,000 images) using the 3-D maximum likelihood (ML3D) method. (a) Comparison between two volumes (#1 [yellow mesh] and #2 [blue surface]) showing absence of bridges (B1b, B8, and BX) in volume #2. (b) Comparison between volumes #1 [yellow mesh] and #3 [white surface], showing absence of bridges (B8 and BX) in volume #3 and presence of additional bridge B1a in volume #3.

Flash photolysis: There exist chemical reagents that become able to react only upon irradiation with light. We have mounted a light pipe leading from a xenon lamp, trained on a point in the path of the grid as it is cryofixed (Shaikh et al. 2009). The two reactants of interests -- one in a unreactive, "caged" form -- are both on the grid at the start of the experiment. The grid is blotted (before the reaction has started) and then flashed. The length of time between flashing and cryo-fixation can be adjusted by the user.

Concurrent with these sample-preparation methods are computational techniques to distinguish between what are likely to be heterogeneous states appearing in images. We have several in-house projects to test the above methods, including the ryanodine receptor with calcium, the ribosome release complex with release factor-1, and various ribosomal complexes with GTP (Shaikh et al. 2010).