Children’s Leukemia Research Award to Fund Myosin Research

(from left to right) Director of Rosenstiel Center Jim Haber, Professor Carolyn Cohen, Dr. Jerry Brown, Anthony Pasqua, President of the Childrens Leukemia Research Association

On April 24, a Children’s Leukemia Research Association (CLRA) award was presented to Jerry Brown, a Senior Research Scientist who works with Carolyn Cohen at the Rosenstiel Basic Medical Sciences Research Center. The award will help fund research on structures of α-helical coiled-coils, in particular those from myosin implicated in certain leukemias. The α-helical coiled coil is a common dimerization motif in proteins and is implicated in many normal physiological as well as pathological processes. Many cases of acute myeloid leukemia involve the aberrant fusion of the transcription factor, CBFβ, to a long portion of the smooth muscle myosin rod, which is predicted from its amino acid sequence to form an α-helical coiled coil. A major aim of the proposed research is thus to crystallize and determine the atomic structures of the segment of the myosin rod nearest this fusion point, both in its normal unfused physiological state and when aberrantly fused to CBFβ. A related aim of the research is to understand how the conformations of α-helical coiled coils in general are affected by attached structures. Accomplishment of these aims may provide a structural basis for the rational design of drugs that can selectively disrupt the activity of the pathologically fused protein.

In addition to Dr. Brown and Professor Cohen, the award presentation was attended by their laboratory researchers Senthil Kumar, Ludmila Reshetnikova, and Elizabeth O’Neall-Hennessey, Rosenstiel Director James Haber, Brandeis Office of Research Administration Associate Director Patricia McDonough, Rosenstiel Department Operations Manager Anahid Keshgerian, CLRA President Anthony Pasqua, his daughter Susan (Pasqua) Bogue, a survivor of leukemia, and Nancy Golden and three of her children.   The award is named after another daughter of Nancy Golden, Amy Golden Uleis, who lost her battle with cancer at age 52 and was a graduate of Brandeis. The award presentation was accompanied by a photo-op and a small reception held at Rosenstiel.

Protein Flexing: A New Look at Transcription-Coupled DNA Repair

Alexandra M. Deaconescu, a research associate in the Rosenstiel Basic Medical Sciences Research Center and 2008-2010 Fellow of the Damon Runyon Cancer Research Foundation, together with Professor of Biochemistry and HHMI Investigator Nikolaus Grigorieff and collaborators in the Laboratory of Dr. Irina Artsimovitch at Ohio State University have just published a new study in PNAS, which delineates novel mechanistic details of transcription-coupled DNA repair.

In any cell, there is intense interplay between various DNA-based transactions, such as replication, transcription and DNA repair. More than twenty years ago, it was discovered that DNA lesions that cause stalling of RNA polymerase molecules elicit a form of preferential nucleotide excision repair (NER) that exists in both eubacteria and eukaryotes, and specifically targets the transcribed DNA strand. Termed transcription-coupled DNA repair (TCR), the process is found to be carried out in bacteria by an ATPase called Mfd or TRCF (see Figure, right). In TCR, TRCF performs two functions: 1) it recognizes a damage-stalled RNA polymerase (RNAP), then dissociates it off the DNA using energy derived from ATP hydrolysis and 2) it recruits DNA repair enzymes via binding to the UvrA subunit of the Uvr(A)BC NER machinery [1].   The Uvr(A)BC machinery is one of the main players in bacterial DNA repair, and distinguishes itself from other DNA repair proteins by its ability to repair a remarkably diverse repertoire of lesions by utilizing a “cut and patch” mechanism, whereby an oligonucleotide containing the damage is excised and the gap later filled.

The cellular role of TRCFs extends beyond TCR. Because of their ability to forward translocate and dissociate stalled RNAPs (or  “backtracked” RNAPs that have slid backwards on the template) [2], TRCFs are also involved in transcription elongation regulation [3, 4], resolution of head-on collisions of the transcription apparatus with the DNA replication machinery [5], and antibiotic resistance [6, 7]. In humans, the effects of impaired TCR are systemic and complex. Mutations in the transcription-repair coupling factor CSB lead to Cockayne Syndrome [8], a progeroid (accelerated-aging) disease characterized by severe developmental abnormalities and neurodegeneration, and whose etiology is currently poorly understood.

To elucidate the mechanism underpinning UvrA recruitment by TRCF, Deaconescu crystallized and solved the X-ray structure of a core UvrA-TRCF complex (Figure, left) demonstrating that UvrA binding involves unmasking of a conserved intramolecular surface within TRCF via a gating motion of the C-terminal domain (red in Figure above). Despite significant effort so far, Deaconescu is still trying to coax nucleotide-bound TRCF to form crystals suitable for X-ray diffraction. These would be highly informative because ATP is required for DNA binding, and its hydrolysis leads to TRCF translocation on dsDNA and ultimately release of RNAP off the damaged template.  Because diffracting crystals eluded her, and to further find out how ADP/ATP modulate the structure of TRCF, Deaconescu learned small-angle X-ray scattering techniques suitable for probing TRCF in solution in the absence and presence of nucleotides, thus circumventing the need for highly-ordered crystals. Then, the Brandeis team and their collaborators at Ohio State employed domain-locking disulfide engineering in conjunction with functional assays to gain a deeper understanding of what TRCF looks like during its catalytic cycle and upon binding to UvrA.  They find that the two main functions of TRCF (RNAP release and UvrA binding) can be uncoupled, suggesting that UvrA recruitment may only occur during/post RNAP release, and not upon RNAP binding as had been proposed earlier in the literature [9]. Furthermore, they show that the ternary elongation complex (consisting of RNAP, template and nascent RNA), but not naked DNA, significantly stimulates ATP hydrolysis by TRCF. Thus, bacterial TRCF operates in a manner reminiscent of that utilized by eukaryotic chromatin remodeling factors, and are preferentially stimulated by nucleosomes over naked DNA substrates.

Deaconescu previously “looked” at TCR using X-rays – as a graduate student she solved the first structure of an intact transcription-repair coupling factor from any organism using X-ray crystallography [10]. She now hopes to reconstitute the larger intermediates that form during TCR and bridge low- with high-resolution information using hybrid structural methods, particularly electron cryo-microscopy, and ultimately formulate a cogent model of how TRCFs operate in cells.

2012 Rosenstiel Award Recipient, Dr. Nahum Sonenberg

2012 Rosenstiel Award Lecture
Thursday, March 29, 2012, 4:00 PM
Gerstanzang 123

The 2012 Rosenstiel award winner, Dr. Nahum Sonenberg of McGill University, is a well-deserving recipient of this honor. Dr. Sonenberg received his Ph.D. in 1976 at the Weizman Institute of Science.  He then worked with Aaron Shatkin, where he discovered the translation initiation factor responsible for binding the 5’ cap of mRNA, eukaryotic Initiation Factor 4E (eIF4E); He has studied translation ever since.  Although his lab focuses on understanding how the cell achieves precise control of translation initiation, this line of investigation has led to discoveries affecting a wide variety of systems.  His lab has made key discoveries in cancer, obesity, virology, memory consolidation and how translation control plays a role in regulating these disparate processes.

In 1988, the Sonenberg lab made the groundbreaking discovery (Nature 1988, http://www.ncbi.nlm.nih.gov/pubmed/2839775) that the uncapped viral mRNA from poliovirus recruits the ribosome to internal regions of the 5’ untranslated region (UTR).  These sites have since been renamed internal ribosomal entry sites (IRESs). This finding was exciting since eukaryotic translation initiation typically requires the 5’ cap on an mRNA for eIF4E binding which subsequently recruits translation initiation machinery.  Until this time, the only mechanism of translation initiation was through the binding of eIF4E to the 5’ cap of mRNAs.  Sonenberg’s discovery that some mRNA has a mechanism to bypass the need for eIF4E binding and thereby avoiding translation control mechanisms started a new line of investigation in the translation field.  Along with discovering IRESs, this paper established an in vitro and an in vivo assay to study cap-independent translation initiation.  These assays are still used widely to test for IRES activity of mRNA UTRs.

Since that initial discovery, it has been found that many viruses contain IRES sequences in the UTR of mRNA that direct translation of viral proteins.  Some viruses, including poliovirus, are able to hijack eukaryotic translation machinery by cleaving factors necessary for canonical cap-dependent translation initiation, but dispensable for IRES translation. In this way, viral mRNAs are able to outcompete eukaryotic mRNAs for ribosome binding and in many cases become the most abundant transcript being translated.

Since the discovery of viral IRESs, many labs, including the Sonenberg lab, have discovered that some cellular genes also use IRESs to bypass the typical translation initiation control mechanisms. These genes are capable of translating even when the cell is actively shutting down translation.  One such cellular IRES-containing mRNA is the insulin receptor message, the IRES I study in the Marr lab.  Using assays similar to those first used in the 1988 paper published by the Sonenberg lab, I am exploring the necessity for the various initiation factors and IRES sequences required for efficient translation of insulin receptor in Drosophila melanogaster and mammalian cells.

The discovery that Dr. Sonenberg made in 1988 is only one example of the elegant research his lab has produced and continues to pursue.

iBiomagazine and iBioseminars

Some video resources if you need to explain scientific topics to students (or need something explained to you!)

iBioMagazine.org features short (<15 min) talks that highlight the human side of research. iBioSeminars.org provides approximately hour-long seminars by high profile researchers.

Professor Emeritus of Biology Hugh Huxley discusses the sliding filament theory of muscle contraction in a November 2011 video from iBiomagazine.org

 

 

Professor of Biology Jim Haber discusses Mechanisms of DNA Repair in a 2009 video from iBioseminars.org

 

James P. Allison to deliver Gabbay Award Lecture

James Allison, PhD  from the Memorial Sloan-Kettering Cancer Center will receive the 2011 Jacob Heskel Gabbay Award in Biotechnology and Medicine “for his development of strategies for the treatment of autoimmune diseases and for immunotherapy of cancer”. The award, administered by the Rosenstiel Center at Brandeis, consists of a $15,000 cash prize and a medallion. Dr. Allison will deliver the award lecture on Mobilizing the immune system to treat cancer: Immune checkpoint blockade, on Monday, Nov 14, 2011 at 3:30 pm in Gerstenzang 121.

Allison and his lab are interested in the mechanisms that regulate T cell responses and using that understanding to improve clinical outcomes in areas ranging from autoimmunity, to allergy to vaccination to  tumor therapy.

Formins require assistance; not so different from other actin nucleators

Formins are a family of proteins conserved across a wide range of eukaryotes and constitute a major class of actin nucleators. In a paper recently published in Molecular Biology of the Cell, a team led by Ph.D. student Brian Graziano in the laboratory of Professor Bruce Goode made the surprising finding that formins depend on co-factors to efficiently nucleate actin assembly both in vitro and in vivo. This discovery was unanticipated because earlier studies had shown that purified formins are sufficient to catalyze actin polymerization in vitro. Graziano, working in collaboration with the labs of Laurent Blanchoin and Isabelle Sagot, investigated the mechanism and function of a formin-binding protein called Bud6 and found that it elevates formin nucleation activity by 5-10 fold. Further, they showed that this activity of Bud6 is critical in vivo for maintaining normal levels of actin cable assembly and polarized cell growth (see figure).

Earlier work from the Goode lab had shown that Bud6 enhances formin-mediated actin assembly in vitro (Moseley et al., 2004), but had left open the question of whether Bud6 stimulates the nucleation or elongation phase of filament growth (an important mechanistic distinction), and whether the activities of Bud6 are important in vivo. Graziano and collaborators dissected Bud6 mechanism by: (a) generating mutations in Bud6 that separately disrupt its interactions with formins (bu6-35) and actin monomers (bud6-8), (b) using TIRF (total internal reflection fluorescence) microscopy to visualize the effects of Bud6 and formins on individual actin filaments polymerizing in real time, and (c) performing a genetic analysis of bud6 alleles. They made three important observations. First, Bud6 enhances the nucleation rather than elongation phase of actin assembly, in sharp contrast to another formin ligand, profilin, that enhances elongation. Second, this activity of Bud6 requires its direct interactions with both the formin and actin monomers, suggesting that Bud6 recruits monomers to the formin to help assemble an actin ‘seed’. Third, genetic perturbation of these activities of Bud6 results in reduced levels of actin cable formation in vivo, in turn causing defects in polarized secretion and cell growth.

Until now, formins were thought to nucleate actin assembly by themselves, which is mechanistically distinct from the Arp2/3 complex (another major actin nucleator). Efficient nucleation by Arp2/3 requires the addition of a nucleation-promoting factor (NPF) such as WASp or WAVE, which recruits actin monomers. Graziano et al. reveal that some formins are similar to Arp2/3 in that they too require an NPF for robust nucleation. Their findings also uncover unanticipated mechanistic parallels between the two systems, since in each case nucleation requires both an actin filament end-capping component (formin or Arp2/3) and an actin monomer-recruiting factor (Bud6 or WASp).

How well is this formin-NPF mechanism conserved? Clues to this question have recently emerged from other studies. A paper published last year in The Journal of Cell Biology by the Goode lab, working in collaboration with the labs of Niko Grigorieff (Brandeis) and Gregg Gundersen (Columbia), implicates the human tumor suppressor protein Adenomatous polyposis coli (APC) in functioning as a formin NPF (Okada et al., 2010). Another study published in The Proceedings of the National Academy of Sciences by the labs of Mike Eck (Dana Farber Cancer Institute), Margot Quinlan (UCLA), and Avital Rodal (Brandeis), suggests that Spire, which is conserved in mammals and flies, may serve as a formin NPF (Vizcarra et al., 2011). Bud6, Spire, and APC all bind multiple actin monomers and interact with the C-terminus of formins to enhance actin assembly, suggesting that they may have related mechanisms and perform functionally analogous roles.

Although the requirement of NPFs increases the complexity of the formin mechanism, it offers an explanation for how cells simultaneously overcome two prominent barriers to actin assembly found in vivo – actin monomer binding proteins (e.g. profilin) that suppress formation of an actin nucleus and capping proteins that terminate growth by associating with the growing end of the filament. NPFs can facilitate nucleation by recruiting actin monomers in the presence of profilin, and formins protect growing ends of filaments from capping proteins. Future work will focus on identifying new formin-NFP pairs, defining the cellular processes with which they are associated, and distinguishing the underlying mechanistic differences among each set.

Cryo-electron tomography and the structure of doublet microtubules

In a new paper in PNAS entitled “Cryo-electron tomography reveals conserved features of doublet microtubules“, Assistant Professor of Biology Daniela Nicastro and coworkers describe in striking new detail the structure and organization of the doublet microtubules (DMTs), the most conserved feature of eukaryotic cilia and flagella.

Cilia and flagella are thin, hair-like appendages on the surface of most animal and lower plant cells, which use these organelles to move, and to sense the environment. Defects in cilia and flagella are known to cause disease and developmental disorders, including polycystic kidney disease, respiratory disease, and neurological disorders. An essential feature of these organelles is the presence of nine outer DMTs (hollow protein tubes) that form the cylindrical core of the structure known as the axoneme. The doublet microtubule is formed by tubulin protofilaments and other structural proteins, which provide a scaffold for the attachment of dynein motors (that drive ciliary and flagellar motility) and regulatory components in a highly specific and ordered manner.

To address long-standing questions and controversies about the assembly, stability, and detailed structure of DMTs , the Nicastro lab used a high-resolution imaging technique, cryo-electron microscope tomography (cryo-ET), to probe the structure of DMTs from Chlamydomonas (single-celled algae) and sea urchin sperm flagella. Cryo-ET involves:

  1. rapid freezing of the sample to cryo-immobilize the molecules without forming ice crystals,
  2. tilting the specimen in the electron microscope to collect ~70 different views from +65° to –65°,
  3. computational alignment of the views to calculate a tomogram (a three-dimensional reconstruction of the imaged sample), and
  4. computational averaging of repeating structures in the tomogram to reduce noise and increase resolution.

Cryo-ET provided the necessary resolution to show that the B-tubules of DMTs are composed of 10 protofilaments, not 11, and that the inner and outer junctions between the A- and B-tubules are fundamentally different (see figure). The outer junction, crucial for the initial formation of the DMT, appears to be formed by interactions between the tubulin subunits of three protofilaments with unusual tubulin interfaces, but one of these protofilaments does not fit with the conventionally accepted orientation for tubulin protofilaments. This outer junction is important physiologically, as shown by mutations affecting the usual pattern of posttranslational modifications of tubulin. In contrast, the inner junction is not formed by direct interactions between tubulin protofilaments. Instead, a ladder-like structure that is clearly thinner than tubulin connects protofilaments of the A- and B-tubules.

The level of detail also allowed the Nicastro lab to show that the recently discovered microtubule inner proteins (MIPs) located within the A- and B-tubules are more complex than previously thought. MIPs 1 and 2 are both composed of alternating small and large subunits recurring every 16 and/or 48 nm along the inner A-tubule wall. MIP 3 forms small protein arches connecting the two B-tubule protofilaments closest to the inner junction, but does not form the inner junction itself. MIP 4 is associated with the inner surface of the A-tubule along the partition protofilaments, i.e., the five protofilaments of the A-tubule bounded by the two junctions with the B-tubule.

The Nicastro lab plans to build on this foundation in future work on the molecular assembly and stability of the doublet microtubule and axoneme, and hope to use it to elucidate molecular mechanisms of ciliary and flagellar motility and signal transduction in normal and disease states.

Other authors on the paper include Brandeis postdocs Xiaofeng Fu and Thomas Heuser, Brandeis undergrad Alan Tso (’10), and collaborators Mary Porter and Richard Linck from the University of Minnesota.

3D electron microscopy reveals: twin spokes are not twins

Movement of cells has fascinated scientists for centuries. Improved handcrafted light microscopes in the late 17th century allowed observations of contracting muscle fibers, single-cell organisms gliding through water drops or cells crawling across surfaces. How cell motility is generated and regulated is an ongoing question researchers at Brandeis and many other institutions are trying to answer. The single-cell green algae Chlamydomonas reinhardtii has two eukaryotic flagella (Fig. A) and is a popular genetic model system for studying these motile organelles, which are also called cilia or undulipodia. Cilia and flagella are basically the same organelles that are highly similar from single-cell algae to humans, but when a cell has many relatively short and asymmetrically beating ones they are called cilia (e.g. on the multi-ciliated epithelial cells that line our airways and are important for mucus-clearance), while a few long ones with often symmetric waveforms are called (eukaryotic) flagella (e.g. the sperm flagellum). These should not be confused with bacterial and archaeal flagella, which are very different in structure and evolutionary origin. Eukaryotic cilia and flagella consist of a microtubule-based, cylindrical core with hundreds of similar building blocks that repeat along the length of the organelle (Fig. B-D). In a single flagellum the activity of thousands of motor proteins, dyneins, has to be coordinated to generate motility, and important regulatory complexes include the radial spokes, in Chlamydomonas two spokes per building block (RS1 and RS2) (Fig. D). Recently, Dr. Thomas Heuser, a postdoc in Dr. Daniela Nicastro’s lab at Brandeis, successfully used three-dimensional electron microscopy (electron tomography) to study the structure of rapidly frozen Chlamydomonas flagella in unprecedented detail (Heuser et al. 2009).

Erin Dymek from Dr. Elizabeth Smith’s laboratory at Dartmouth College found that the concentration of Calcium ions, a known regulatory signal modulating ciliary and flagellar motility, affects dynein activity through a conserved Calmodulin and Radial Spokes associated Complex (CSC) (Dymek and Smith, 2007). Erin Dymek and Elizabeth Smith have now teamed up with Tom Heuser and Daniela Nicastro to study the 3D location of this Calcium sensing complex in flagella. In a recent paper (Dymek et al. 2011 MBoC in press) they compared the wild type structure of Chlamydomonas flagella to several artificial microRNA-interference mutants lacking parts of the CSC. They found that in all amiRNAi mutants many of the flagellar building blocks were missing one specific radial spoke, RS2, while RS1 was always present (Fig. E-G), suggesting that the Calcium sensing CSC is located at or near RS2. Interestingly, RS1 and RS2 were previously assumed to be structurally identical, their different numbering simply referred to their proximal and distal location within the repeating building block. The current study not only indicates that the CSC is required for spoke assembly and wild type motility, but as one of the most surprising outcomes it also provides evidence for heterogeneity among the radial spokes, at least at the base where the spokes are anchored to the microtubules. The same team of biologists is now continuing to study the CSC location in the flagellar building block in greater detail by improving image processing strategies to increase resolution.

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