Molecules that Walk

The Department of Biological Science’s contribution to Science week 2016 kicked off on 11 April with a lecture by Dr Anthony Roberts, a young Principal Investigator who arrived at Birkbeck in 2014. Anthony received his B.Sc. from Imperial College in London and his Ph.D. from the University of Leeds, and spent four years as a postdoc at Harvard in the USA before moving here to start his own research group as a Sir Henry Dale Fellow of the Wellcome Trust and Royal Society.

Anthony began his lecture by explaining that he was going to talk about molecules that have the capacity to produce directed movement – or to ‘walk’ – and their importance for human health. These molecules are all proteins, and the context in which they move is the interior of living cells. Both the proteins he studies, kinesin and dynein, ‘walk’ on a network of highways conceptually not unlike the transport system that we use to move around London. These cellular highways are filaments called microtubules, which, unlike our roads and railway tracks, are able to self-assemble and also to self-destruct.

The ability to move is one of the fundamental properties of life, and scientists and philosophers have been studying it for millennia. Muscles were identified as the organs of movement in antiquity, but it was not until the mid-twentieth century that the molecules involved in muscle contraction could be identified. The Hungarian physiologist Albert Szent-Györgyi discovered the muscle proteins now named actin and myosin using very simple equipment during the Second World War.

These proteins have similarities with kinesin and dynein, although historically they have been easier to study due to their abundance in muscle; actin forms fibrils and the enzyme myosin binds to and ‘walks’ along these filaments. This process, like all movement, requires energy, and this is obtained from the cell’s power source, the small molecule adenosine triphosphate (ATP). The part of the myosin molecule that binds to actin, which is called its head, breaks a phosphate bond in this molecule to liberate energy and power the walking motion; many of these ‘power strokes’ together cause the muscle fibre to contract.

Ideally, we would want to watch this, or any other form of molecular motion, in real time, but this is impossible because molecules are far too small: smaller than the wavelength of light, so they cannot be viewed in a light microscope. Studies of molecular structure require techniques like X-ray crystallography and electron microscopy, both of which have been used to study motor molecules.

However, neither of these techniques can do more than generate still images. Movement can only be inferred by taking lots of snapshots of the molecules at different points during the movement cycle, rather like the earliest movies. We have now built up a complete picture of actin and myosin that is detailed enough for the positions of individual atoms to be seen clearly.

Not all movement in nature, however, uses muscles. Single-celled organisms – the ‘animalcules’ observed by pioneer microscopist Antonie van Leeuwenhoek in the 1670s – have directed movement, as do bacteria, and these have neither muscles nor nervous systems. And directed movement also occurs inside cells. A good example of this is the division of replicated DNA between daughter cells during cell division.

The interior of all cells is a viscous mixture, crowded with molecules; it is possible for small molecules to move from one part of a cell to another through diffusion, but this process would be impossibly slow for larger ones. Motor proteins, on the other hand, can carry ‘cargo’ molecules across cells remarkably quickly and efficiently. Motor proteins can traverse a distance of 0.1 mm – the length of a large animal cell – in two minutes, which in terms of lengths per second is approximately three times faster than a car.

Both the motor proteins studied in Anthony’s lab, kinesin and dynein, ‘walk’ along microtubules inside cells. These filaments typically form with one end towards the centre of the cell, and its nucleus, and the other towards the cell periphery, and the motor proteins move in opposite directions: dynein towards the nucleus, and kinesin towards the cell edge.

Any kind of directed movement by molecules is challenging for several reasons. Motor molecules have no equivalent of our nervous systems for controlling movement, and they are far too small to be held on their tracks by gravity; instead, they grip the microtubules using chemical forces. They experience negligible inertia, and are constantly buffeted by other molecules in the cell. It would therefore be catastrophic for the whole of a walking molecule to leave its path at once.

The structure and function of conventional kinesin are now fairly well understood. It consists of two identical protein chains, and each chain has two major domains separated by a short linker. The larger domain of each chain coils together to form a single long stalk; the smaller domain is globular and attaches to the microtubule, so the molecule looks rather like a single leg with two feet. Each of the feet is an enzyme that generates the energy for the motion by breaking down ATP to form ADP and release a phosphate group, and it cycles between ATP-bound, ADP-bound and empty states.

The step between ADP-bound and empty is a bottleneck that can be relieved when the foot attaches to the microtubule in a particular position, ensuring that the whole molecule moves in the correct direction. The trailing foot is released from the microtubule and the cycle begins again once ATP has bound to the front foot, triggering a conformational change in the whole molecule.

The core of kinesin is similar in structure to myosin, suggesting that these two proteins have a common ancestor. The other microtubule-bound motor protein, dynein, has a different origin. Although we still know comparatively little about it, it was actually the first of the microtubule-bound motor proteins to be discovered: this was in the 1960s, when it was found as the protein that generates the force that allows protozoa and sperm cells to swim. Anthony’s group, however, has been studying how it functions inside cells to move ‘cargo’ – often nucleic acids or other proteins – from the edges of the cell towards its interior. It also helps to pull the duplicated genetic material between the two halves of the cell during cell division.

Dynein is a much larger and more complex molecule than the other motor proteins. Its structure, like those other proteins, has several components: in this case, a stalk, a ring and a tail, with a linker between the stalk and the ring. Much of what we know about this large structure has come from electron microscopy, and more recently X-ray crystallography.

The structure of dynein; the stalk is shown in yellow and the linker in magenta.

Anthony’s group and others have developed a model in which the main mechanical element is the linker, which bends and straightens to displace the cargo-bound end of the structure along the microtubule in the direction of travel. The image shown here is a still from an animated model of how dynein generates movement, which remains speculative in places and is helping to stimulate new experiments in these areas. It is also incomplete, as it only shows one half of the molecule: we do know that dynein, like kinesin, is a biped, but exactly how its ‘feet’ are coordinated remains at the frontier of our knowledge.

Anthony ended his talk by discussing some actual and potential medical applications of studies of walking molecules. Some commonly used anti-cancer drugs, including taxol, work by stabilising microtubules to prevent motion and therefore stop cancer cells from dividing. Molecules that interact with motor proteins are also being studied as potential treatments for neurodegenerative diseases and for some types of heart disease. One such compound is a myosin activator, omecamtiv mecarbil, which is showing promise as a treatment for heart failure. And we are likely to discover further applications as we learn more about these fascinating walking molecules.

Crystallography: from Chocolate to Drug Discovery

Birkbeck has already established lecture series in honour of some of its most distinguished alumni. Until 2016, however, Rosalind Franklin – co-discoverer of the DNA structure and perhaps the most widely recognisable of its ‘famous names’ – was missing from the list of honourees. This gap has now been filled; the annual Rosalind Franklin lecture forms part of the college’s Athena SWAN programme and will always be given by a distinguished woman scientist. And fittingly, the inaugural lecture, which was part of Science Week 2016, was devoted to Rosalind Franklin’s own discipline, crystallography. Elspeth Garman, Professor of Molecular Biophysics at Oxford University, gave an entertaining and illuminating lecture to a large audience that included Rosalind’s sister, the author Jenifer Glynn.

Garman began her lecture by showing a short video that she had produced for OxfordSparks.net that used a ‘little green man’ to illustrate the method of X-ray crystallography that is used to obtain molecular structures from crystals. The rest of the lecture, she said, would simply go through that process more slowly. She started by showing some beautiful examples of crystals. All crystals are formed from ordered arrays of molecules. They can be enormous, such as crystals of the mineral selenite in a cave in Mexico that measure over 30’ long or too small to be visible with the naked eye.

In the early decades of crystallography, structures could only be obtained from crystals of the smallest, simplest molecules: the first structure of all, published in 1913 by the father-and-son team of W.H. and W.L. Bragg, was of table salt. When they were jointly awarded the Nobel Prize for Physics in 1915, the younger Bragg was a 25-year-old officer in the trenches on the Western Front. His record as the youngest Nobel Laureate was unbroken until Malala Yousafzai’s Peace Prize in 2014.

The Braggs’ discoveries paved the way for studies of the structures of many, many substances: including the chocolate of the lecture title. Few of the audience can have known that chocolate exists in six different crystal forms, or that only one of these (Form V) is good to eat. The process of ‘tempering’ – a series of heating and cooling steps – is used to ensure that it solidifies in the correct form.

Garman then moved on to talk about her own field of protein crystallography. Proteins are the ‘active’ molecules in physiology, and they are formed from long, linear strings of 20 different ‘beads’ (actually, small organic molecules known as amino acids). Chemists can quite easily find out the sequence of these beads in a protein, but it is impossible to work out from this the way that the string will fold up into a definite structure ‘like a piece of wet spaghetti’. And it is this structure that places different units with different chemical properties on the surface or in the interior of the protein, or near each other, and that therefore determines what the protein will do.

Protein crystallography only became technically possible in the mid-twentieth century, and even then it was a painfully slow and complex process that could only be used to study the smallest, simplest proteins. Dorothy Hodgkin, also a professor at Oxford, won her Nobel Prize in Chemistry in 1964 for the structures of two biologically important but fairly small molecules: penicillin, with 25 non-hydrogen atoms and vitamin B12, with 80. She is perhaps better known for solving the structure of insulin, the protein that is missing or malfunctioning in diabetics. This has 829 non-hydrogen atoms; in contrast, the 2009 Chemistry Nobel Prize was awarded for the structure of the ribosome, the large (by molecular standards) ‘molecular machine’ that synthesises proteins from a nucleic acid template. The bacterial ribosome used for the Nobel-winning structural studies is well over 300 times larger than insulin, with over a quarter of a million atoms.

Protein structures are not only beautiful to look at and fascinating to study, but they can be useful, particularly for drug discovery. Many useful drugs have already been designed at least partly by looking at a protein structure and working out the kinds of molecule that would bind tightly to it, perhaps blocking its activity. Some viral proteins have been particularly amenable to this approach. Rosalind Franklin did some of the first research into virus structure when she was based at Birkbeck, towards the end of her tragically short life, and her student Aaron Klug cited her inspiration in his own Nobel lecture in 1982. X-ray crystal structures were used in the design of the anti-flu drugs Relenza™ and Tamiflu™ and of HIV protease inhibitors, and more recently still structures of the foot and mouth virus are helping scientists develop new vaccines for tackling this potentially devastating animal disease. The foot-and-mouth virus structure even made the front page of the Daily Express.

The equipment that Dorothy Hodgkin and her contemporaries used to solve protein structures in the 1960s and 1970s looks primitive today. Now, almost every step of protein crystallography has been automated. Powerful beams of X-rays generated by synchrotron radiation sources, such as the UK’s Diamond Light Source in Oxfordshire, allow structures to be determined quickly from the smallest crystals. It is even possible to control some of these machines remotely; Garman has operated the one at Grenoble from her sitting room. Yet there is one step that has changed remarkably little. It is still almost as difficult to get proteins to crystallise as it was in the early decades. Researchers have to select which of a large number of combinations of conditions (temperature, pH and many others) will persuade a protein to form viable crystals. Guesswork still plays a large part and some researchers seem to be ‘better’ at this than others: Garman adds the acronym ‘GMN’ or ‘Grandmother’s maiden name’ to her list of conditions to reflect this.

Yet, with every step other than crystallisation speeded up and automated beyond recognition, the trickle of new structures in the 70s and even 80s has become a torrent. Publicly available structures are stored online in the Protein Data Bank, which started in 1976 with about a dozen structures: it now (May 2016) holds over 118,000. Protein crystallography as a discipline is thriving, but there are many challenges ahead. We are only now beginning to tackle the 70% or so of human proteins that are only stable when embedded in fatty cell membranes and are therefore insoluble in water. It is possible to imagine a time when it is possible to solve the structure of a single molecule, with no more need for time-consuming crystallisation. And, hopefully, women scientists will play at least as important a role in the second century of crystallography as they – from Quaker Kathleen Lonsdale, who developed important equations while jailed for conscientious objection during World War II, through Franklin and Hodgkin to Garman and her contemporaries – have in the first.

Evolution and Assembly of Protein Complexes

Since 2003, all Birkbeck researchers in structural biology and allied disciplines have collaborated with colleagues at UCL in the Institute of Structural and Molecular Biology. The Institute holds a varied series of events throughout the year, including a programme of research seminars arranged termly around current themes in molecular and structural biology research. The theme for the spring term 2016 seminar programme has been ‘Protein Dynamics: from Folding to Function’; one of the first of the distinguished scientists invited to present their research under that theme was Sarah Teichmann from the EMBL – European Bioinformatics Institute and the Sanger Institute at Hinxton, near Cambridge, UK. She gave a fascinating talk that linked evolution and protein folding to the topic of Section 7 of the PPS course, quaternary structure (or the assembly of protein complexes).

Teichmann has won many awards in what is still quite a short research career, including the Biochemical Society’s Colworth Medal for ‘an outstanding research biochemist under the age of 35’ (2011) and the EMBO Gold Medal (2015). Last year she was elected a Fellow of the prestigious Academy of Medical Sciences, with a citation that commended her as representing ‘a new breed of scientists at the interface between computational and experimental molecular biology’. She is also an advocate for women in science and has written a children’s novel.

She began her seminar by asking two related questions: ‘how do protein complexes assemble?’ and ‘how do protein complexes evolve?’ and by misquoting the poet John Donne: ‘no protein [man] is an island’. Many proteins are functional only when bound to others to form complexes, and in the crowded environment of a cell each newly synthesised protein has only a limited amount of time to find its partners and form a stable complex. Much can be learned about the evolution and dynamics of complex formation by studying the complexes that are available in the Protein Data Bank. Her group’s evaluation of these structures has contributed to the software that the PDB uses to predict the functional biological unit (monomer, dimer or multimer) for each structure in the PDB, and has led to the 3Dcomplex.org database of protein complexes. This database provides a hierarchical classification of now over 30,000 protein quaternary structures. Each complex is represented using graph theory as a simple 2D figure or ‘mini-graph’, with each polypeptide chain as a node and each interaction surface between two chains as an edge. These little graphs make it easier to distinguish between topologies involving the same number of subunits: for example, a complex of six identical protein chains may be a simple hexamer with 6-fold rotational symmetry (such as the traffic ATPase [PDB 1g6o]) or a dimer of trimers with 32 symmetry (such as annexin XII [PDB 1aei]). The links here are to the pages describing those proteins in Section 7 of the PPS course material.

Alongside the hierarchy described in the 3Dcomplex database, protein complexes can be divided into two large groups: homomers, which consist of multiple copies of the same polypeptide chain, and heteromers involving different chains. (Haemoglobin, a tetramer with two alpha and two closely related beta chains, is arguably an intermediate between the main two types.) Teichmann spent the rest of her lecture addressing three related questions about the assembly of both homomers and heteromers:

i) Does the assembly of protein complexes drive evolution?
ii) What are the mutational mechanisms involved in complex formation?
iii) Can the principles of protein assembly be used to predict topologies that have not yet been seen?

Starting with the first question, from an evolutionary point of view the simplest complex to form is a homodimer with two copies of the same monomer; one mutation that turns part of a protein surface into a ‘sticky patch’ is all that is necessary to stabilise dimer formation. Not surprisingly, the homodimer is also the commonest type of quaternary structure found in the PDB. Once a protein has dimerised, additional monomers can be added to form larger complexes with cyclic symmetry, or the dimer itself can (for example) dimerise. The order in which the interfaces in a multimer formed during evolution can be predicted from the amount of surface area buried by the formation of each interface, with the largest surface areas being buried first. This simple rule applies to complicated assemblies as much as to simple ones, and to heteromers as much as to homomers. Therefore, for all but the very simplest structures, it is almost impossible to predict the form that a complex will take unless you know the order in which the subunits assemble. Joseph Marsh, a former postdoc in Teichmann’s group now working at the MRC Human Genetics Unit in Edinburgh, represents this here in an analogy with the assembly of flat-pack furniture, with and without instructions.

Illustration © Joseph Marsh, MRC Human Genetics Edinburgh

Teichmann tested some of her predictions of protein assembly pathways using mass spectrometry in collaboration with Professor Dame Carol Robinson’s group at the University of Oxford, and found that seven out of nine pathways and 22 out of 27 steps within those pathways had been predicted correctly. This hierarchy of subunit assembly can also be used to predict the evolution of a complex, so it is clear that the assembly of protein complexes can indeed drive evolution.

Turning to the second question, Teichmann used specific examples of protein families that take up different quaternary structures in different species, including the PyR family of bacterial pyrimidine operon attenuators, to explore the evolutionary mechanisms that take a protein from one that is most stable as a monomer to different multimeric forms. These can involve direct mutations at the interface between subunits (for example, making the protein surface ‘stickier’ or creating a salt bridge) and other so-called ‘allosteric’ mutations that change the protein structure to allow different interfaces to form. Often, the difference between (for example) a protein that is stable as a dimer and one that is stable as a tetramer will come down to changes in a few amino acids. In the case of the PyR attenuator family, mutations away from the interface drive a conformational change that is equivalent to the one that occurs when the protein binds DNA, and so stabilise multimer formation.

Finally, Teichmann considered the use of the assembly principles that she had outlined in predicting the form that a protein complex would take from scratch. Most basic steps in complex assembly, as described earlier, can be grouped into one of three categories: dimerization of one or more chains, adding an identical subunit or subunits to a complex (cyclization) and adding a different type of subunit. These can be combined in different ways to form a large number of possible quaternary structure topologies. So far, about 120 different topologies are represented in the PDB, with four or five new ones being added each year, and the vast majority of these fit into one of Teichmann’s topologies. She assembled all the predicted topologies, including those not yet observed, into a ‘periodic table of protein complexes’ (S.E. Ahnert et al., Science 350, aaa2245 (2015)). This table has already been seen to correctly predict the topology of some newly determined complexes that were not included in the original list.

Bernal Lecture 2015: Terminating Protein Synthesis

Professor J.D. Bernal, known to his colleagues and contemporaries as ‘Sage’, spent much of his career as a professor of Physics at Birkbeck and became the first chair of the Department of Crystallography in 1963. When he retired in 1968 the college founded a series of lectures in his honour. The first of the Bernal Lectures, in 1969, was given by Dorothy Hodgkin, winner of the 1964 Nobel Prize for Chemistry for solving the structures of ‘important biological substances’, mainly penicillin and vitamin B12. In 2015 we were honoured to welcome another Chemistry Nobel Laureate to give this annual lecture. Professor Sir Venki Ramakrishnan of the MRC Laboratory of Molecular Biology in Cambridge was awarded the prize in 2009 with Thomas Steitz of Yale University, USA and Ada Yonath of the Weizmann Institute of Science, Israel, for studies of the structure and function of the ribosome: the ‘molecular machine’ that catalyses the synthesis of proteins from their messenger RNA (mRNA) templates.

The lecture, held on 19 October 2015, was introduced by David Latchman, Master of Birkbeck College and Professor of Genetics. He welcomed three generations of Bernal’s descendants to Birkbeck, highlighted the success of the lecture series in attracting some of the most distinguished researchers in structural biology and allied disciplines, and explained that the topic of the lecture overlapped with some of his own research interests in the regulation of gene expression.

Prof. Ramakrishnan began his lecture by explaining that protein synthesis was a complex process, involving many proteins as well as the ribosome itself, and that he would be talking about a particular point in this process: namely, how it ends (formally, the termination of protein translation). He showed an image of a ribosome in the process of protein synthesis that, he explained, represented the culmination of 40 years’ work on its structure, and explained how the linear mRNA molecule wound through a cleft between the two subunits of the ribosome. As the mRNA passes through the ribosome each of its three-base ‘codons’ comes into contact with three sub-parts of the ribosome’s active site – the A-site, P-site and E-site – in turn. When a codon enters the A-site it binds to the anti-codon of the transfer RNA (tRNA) carrying the next amino acid; the amino acid is bonded to the previous amino acid in the growing protein chain in the P-site, and the now empty tRNA released from the ribosome in the E-site. This continues until the new protein chain is complete. This is signalled by one of the so-called ‘stop codons’ UGA, UAG and UAA, which have no corresponding tRNAs, entering the ribosome’s A-site. The new protein is released from the ribosome to fold into its native structure, and the ribosome subunits dissociate.

Diagram of a ribosome showing the three tRNA binding sites during protein elongation
Taken from the PDB ‘Structural View of Biology: The Ribosome’
© David Goodsell, 2010

Although the process of adding amino acids to a protein chain is extremely similar in all organisms, there are significant differences between bacteria (prokaryotes) and eukaryotes in the process of termination – as, indeed, there are in the initiation of protein synthesis. We are beginning to understand these mechanisms only now that we can obtain high resolution structures of ‘snapshots’ of the ribosome at different points during the protein synthesis cycle and follow the sequence of conformational changes that occur then.

All stop codons are recognised and decoded by proteins known as a release factors. Bacteria have two of these: RF1 recognises UAG, RF2 recognises UGA and they both recognise UAA. Eukaryotes have only one RF, which can recognise all these codons. These three proteins all have a common sequence motif, GGQ, which is known to be involved in the release of the protein from the ribosome. The structures of the eukaryotic and prokaryotic release factors are different, but all bind to the ribosome in such a way that the GGQ motif and the part of the structure that recognises the stop codon are exactly the same distance apart as the length of a tRNA molecule. These parts of the protein will therefore interact with the peptide and the stop codon at the same time.

Prof. Ramakrishnan and his group spent years trying to obtain near atomic resolution structures of functional ribosome-release factor complexes; this problem was solved initially for the smaller prokaryotic ribosomes but now for eukaryotic ones as well. In all cases, the release factors bind to the ribosome in a different way to the tRNA molecules, inducing a different conformational change in the complex. Using the bacterial structures, the group was able to understand why the factors RF1 and RF2 only recognise the codons that they do.

In the case of eukaryotes, not only were the structures harder to obtain, but the basic question to be asked was more complex: how can a single protein recognise the stop codons UGA, UAG and UAA, but not UGG (which codes for the amino acid tryptophan)? Ramakrishnan reasoned that mutating the GGQ motif in the release factor would make it inactive, and that binding this mutant protein to the ribosome with an ATPase might ‘trap’ the complex in the structure that it takes up before protein is released and allow the structure to be determined. Electron micrographs of these structures have shown that the three anti-codons and no others are recognised through a combination of base stacking and hydrogen bonding. Ramakrishnan ended his talk by comparing anti-codon binding to a NAND gate in electronics, with G representing ‘1’ and A ‘0’: any combination except GG (‘11’) in the second and third anti-codon positions leads to termination of translation and protein release.

This work was published in Nature in August 2015 and the (very large!) structures of the complexes – one snapshot with the release factor bound in each of the ribosome subsites – are available in the PDB as entries 3JAG,3JAH and 3JAI. These are some of the most recent of the 103 PDB structures on which Prof Ramakrishnan has so far been named as an author; you can view them all on a timeline on the PDB site.

Note: If you are reading this blog post as a current PPS student, don’t be surprised if you find it difficult to understand. We will cover the structure and mechanism of the ribosome later in the course (in Section 8: The Protein Lifecycle). If you bookmark this blog post and come back to it after you have studied that section you should find that you can make much more out of it.

Welcome to PPS students 2015-16!

This post is extremely like those I have written at this time of year for the past few years. This is because what I have to say now is very, very similar...

I would like to offer a warm welcome to the Principles of Protein Structure blog to all students who have just started studying Birkbeck's Principles of Protein Structure (PPS) course, and a welcome back to any who have taken a break in studies and intend to complete the course this year.

I run this blog to link the material that you will be studying in the course to new research developments in the areas of protein structure and function and related aspects of biotechnology and medicine. I might, example, report on talks given in the ISMB seminar series run jointly by the Department of Biological Sciences at Birkbeck and research departments in neighbouring University College London. The programme for Autumn 2015 focuses on transcription, which is the process through which DNA is copied into RNA; this will be covered in some detail in one of the later sections of the course, the Protein Lifecycle. Other posts may be reports from conferences or summaries of recently published papers in protein structure, protein bioinformatics and allied areas. Look out for an account of a lecture to be given at Birkbeck in October by Venki Ramakrishnan, who was awarded a share of the Nobel Prize for Chemistry in 2009 for structural studies of ribosomes and will shortly take over as President of the Royal Society.

Some posts on this blog are written by "guest blogger" Jill Faircloth, who took the MSc in Structural Molecular Biology a few years ago and is now working as a freelance science communicator. She introduces herself in this post written in March 2012, in which she also describes how she found the later part of the PPS course and her thoughts on the two choices available for the second year of the MSc.

Do, if you get a chance, look through some blog posts from earlier years to see the kind of topics that we will be discussing. However, don't be discouraged if at this stage of the course you find the science presented there difficult to understand. I can assure you that it will get easier!

I particularly recommend that you look at a couple of posts from December 2013 and July 2014 about the history of structural science, particularly X-ray crystallography. Crystallography was the first method to be developed for solving the structure of biological macromolecules, and it is still the most important. The year 2014 was designated by the United Nations as the International Year of Crystallography, marking the year between the centenaries of the publication of the first papers on X-ray diffraction and the award of the 1915 Nobel Prize for Physics to the father-and-son team of William and Lawrence Bragg who made the principal discoveries.

So - the best of luck for the 2015-16 PPS course and for your studies at Birkbeck! We hope that many of you will go on to complete our MSc in Structural Molecular Biology.

Best wishes,

Dr Clare Sansom Senior Associate Lecturer, Biological Sciences, Birkbeck and Tutor, Principles of Protein Structure

Microtubules and Microscopes: Exploring the Cytoskeleton

Electron microscopist Carolyn Moores, the most recently appointed professor in the Department of Biological Sciences at Birkbeck, gave her inaugural lecture at the college on June 1.

Moores arrived at Birkbeck in 2004 to start her research group and has risen rapidly and steadily up the academic ladder ever since. Introducing the lecture, the Master of Birkbeck, David Latchman, explained that Moores’ CV stood out in every way; she was clearly as gifted a teacher and administrator as she was a researcher. Furthermore, as she has won several awards for science communication, he predicted that the audience would be in for a treat. We were not disappointed.

Moores began her lecture by saying that she would talk about three different things: her own career development; her group’s research into the structure and function of microtubules; and the advancement of women in science, a cause that is close to her heart.

She remembered that she had wanted to work as a scientist as soon as she knew what a laboratory was, and she started young, as an intern in a research lab at Middlesex Hospital while still in the sixth form. School was followed by a BSc in Biochemistry at Oxford and a PhD in John Kendrick-Jones’ lab at the world-famous Laboratory for Molecular Biology (LMB) in Cambridge. She then moved to work as a post-doc with Ron Milligan at the Scripps Research Institute in La Jolla, California, USA, and it was there that she began her studies on microtubules.

The award of a David Phillips research fellowship in 2004 gave her the opportunity to return to the UK as an independent researcher. She explained that there were three reasons – or more accurately three people – that led her to choose to come to Birkbeck. Working in electron microscopy, she was inspired by the work of Helen Saibil, one of the UK’s principal exponents of that technique; she had known Nicholas Keep, then a lecturer in Biological Sciences, as a friend since her time at the LMB; and she knew that she would value the interdisciplinary working environment of the Institute for Structural Molecular Biology under the ‘inspired’ leadership of Gabriel Waksman.

Moores then moved on to discuss the main topic of her group’s research: the three-dimensional structure, function and role in disease of tiny cylindrical structures known as microtubules. These are one of the building blocks of the cytoskeleton, which forms a framework for our cells in the same way that our skeletons form a framework for our bodies. They are about 25nm in diameter, which puts them firmly into the ‘nano-scale’ of biology that is easily studied using electron microscopy.

There is a cytoskeleton in every living cell, and it, and the microtubules that form it, are involved in many important cellular processes including shape definition, movement and cell division. Diseases as diverse as cancer, epilepsy, neurodegeneration and kidney disease have been linked to microtubule defects. Understanding their fundamental structure and function, as Moores’ group aims to, should help in understanding these disease processes and perhaps also in developing effective treatments.

Microtubules are built up from many copies of a small protein called tubulin, which, in turn, is a dimer of two similar proteins called alpha and beta tubulin. These tubulin dimers make contacts with each other both head-to-tail and side-to-side to create the cylindrical microtubule wall, fuelled by energy derived from the molecule GTP. Each tubulin unit has a definite “top” and a “bottom” and, as the units are oriented in parallel, so has the complete microtubule.

Microtubules are dynamic structures; they continue to grow by the addition of tubulin units to one end as long as GTP is available, and then begin to unravel and shrink. This dynamism, which allows them to respond to the changing needs of the cell, is essential for their function in healthy cells. In particular, microtubules organise chromosome structures during cell division and are therefore necessary for cell proliferation. As cancer is a disease of uncontrolled cell proliferation, it is possible to imagine that a molecule that could specifically block microtubule growth and assembly in the nucleus might be useful as an anti-cancer drug.

Moores and her group are aiming to understand the process of microtubule growth at as high resolution as possible, using electron microscopy. Unfortunately, however, the most detailed images can only be obtained if the specimen is at very low temperatures (in so-called cryo-elecron microscopy) and using this means that the dynamics of the specimens must be “frozen” into a still image. While it is now possible to see the individual tubulin subunits in the static microtubule images, many details of their structure can only be inferred from computational analysis.

Moores went on to describe one project in her lab in a little more detail. This was an investigation of the structure and role of proteins that bind only to growing microtubule ends, falling off when the growth stops. It is possible to obtain low-resolution images of microtubules in which these molecules have been made to fluoresce, so only growing microtubules are tracked.

In order to understand the growth process in detail, the group developed an analogue of the GTP “fuel” molecule which can bind to the tip of a microtubule that is extending but not break down to release its energy, so the microtubule does not in fact grow. This forms a static analogue of a growing microtubule that retains all the characteristics of the dynamic structure but that can be studied at low temperatures.

The binding site for End Binding protein 1 (highlighted in green) on the microtubule lattice at the corner of four tubulin dimers, visualised using cryo-electron microscopy
Image from Maurer et al. 2012, Cell 149(2):371-82. Full text here.

Images of this structure have shown that the end binding proteins bind at the corner of four of the tubulin units. They have explained a lot of the properties of growing microtubules, but there is still more to learn. A full understanding will need structures that are at even higher resolution, where the positions of individual atoms can be made out. Following many years of technical development, today’s most powerful electron microscopes are now making this possible.

In the last section of the lecture, Moores left the topic of research to talk briefly about another of her passions: the promotion of women in science. She explained that although 65% of under-graduates in the biological sciences are now women, the proportion of women drops to 40% at any academic grade and 25% for full professors.

A study cited by the European Molecular Biology Organisation has suggested that the barriers for women scientists to progress are set so high that at the current rate of progress full equality would never be achieved. Birkbeck has signed up to the Athena SWAN Charter, set up to encourage higher education institutions to transform their culture and promote gender equality. She described her work with the Athena SWAN team that has so far resulted in the college gaining a bronze award as being as exciting as, but also as challenging as, her studies of microtubules.

There is more information about the structure and function of microtubules on this page in section 7 of the PPS course. The technique of cryo-electron microscopy is covered in some detail in the second-year MSc module Techniques in Structural Molecular Biology.

Protein Machines in the Molecular Arms Race

Birkbeck’s Science Week 2015 was held from Monday 23 to Thursday 26 March and included three evenings of public talks by senior researchers. The first two lectures, on the Tuesday, were given by two of the college’s most distinguished women scientists: Helen Saibil from the Department of Biological Sciences and Karen Hudson-Edwards from Earth and Planetary Sciences; they were billed together as a ‘Women in Science Evening’.

The lectures were all introduced by the Dean of the Faculty of Science, Nicholas Keep who described Saibil, a close colleague, as “our most eminent female scientist”. She came to Birkbeck from Canada via a PhD at King’s College London under the supervision of Nobel laureate Maurice Wilkins and post-doctoral work at Oxford.

Since arriving here in the 1980s she has built up an internationally renowned structural biology lab, focusing in particular on the technique of electron microscopy. She has been a Fellow of the Royal Society since 2006 and of the Academy of Medical Sciences since 2009.

Saibil began her lecture by explaining that proteins can act as little machines, performing mechanical tasks that are essential for the maintenance of life. Her group has been interested for some time in proteins that can punch holes in the walls of cells. This allows the cell contents to leak out in a damaging process known as lysis, and it also allows toxins to enter the cells. These proteins can therefore be thought of as powerful weapons, and they are deployed on both sides of a ‘molecular arms race’: by pathogens and by the immune systems of humans and other animals.

Most soluble proteins fold into a single stable structure that tries, as far as possible, to keep their hydrophobic (“water-hating”) parts – the side chains of certain amino acids – in the interior of the protein, with the hydrophilic (“water-loving”) side chains on the outside, in contact with the watery environment inside or outside cells.

Pore-forming proteins, however, have a ‘Jekyll and Hyde’ like identity: they can form two distinctly different shapes, one as individual, soluble molecules and the other when they associate with each other into membrane-bound rings to form cylindrical pores. These structures, and the conformational change between them, are remarkably similar in proteins from bacteria and from the immune system.

Pore-forming toxins have been found in types of bacteria that are responsible for some deadly human diseases, including meningitis and pneumonia. The structure of a monomeric form of one of these proteins in solution was first solved in 1998, using X-ray crystallography. However, large complexes of many protein molecules are more readily solved by electron microscopy, particularly when those complexes are embedded in membranes.

In 2005 Saibil and her group described structures of the pore-forming toxin pneumolysin from Streptococcus pneumoniae, in complex with a model cell membrane. They found that the proteins formed two distinctly different ring-shaped structures. Initially, they formed into a ring sitting on top of the membrane, which was termed the pre-pore; then they changed shape to burrow part of each protein deep into the membrane and form the pore itself. Each monomer in the pre-pore had a structure that was similar to that of the molecule in solution, but they underwent large structural changes to form the pore.

Schematic illustration of how suilysin, a bacterial cholesterol-dependent cytolysin, drills holes in cell membranes. Image © Adrian Hodel, London Centre for Nanotechnology

Most structures solved by electron microscopy are at lower resolution than those solved by X-ray crystallography, and it is not possible to trace the positions of individual atoms at lower resolutions (e.g. worse than 3 A). Saibil and her colleagues were able to interpret the structure of the proteins making up the pore by fitting pieces of the X-ray structure of the isolated molecule into their electron density.

They found a dramatic change in structure, with the tall, thin protein structure collapsing into an arch and a helical region stretching out to form a long, extended beta hairpin. It is these hairpins that join together to form the walls of the pore. The process of pore forming therefore has three stages: firstly the toxin molecules bind to the surface of their target cells, then they associate into the circular pre-pores and finally they change shape in a concerted manner, punching holes in the cell membranes by ejecting a disc of membrane, letting other toxins in and cell contents out.

Saibil then turned the focus of her talk from attack by bacteria to the human immune system’s defence. Natural killer (NK) cells are specialised lymphocytes (white blood cells) that kill virally-infected and cancerous cells in the bloodstream. They kill on contact with their target cells by releasing a toxic protein into those cells that stimulates those target cells to commit suicide in a process known as programmed cell death or apoptosis. We have only recently learned that the mechanism through which the NK cells work is very similar to the mechanism of the bacterial pore-forming toxins.

Natural killer cells express a protein called perforin that has a similar structure in solution to the bacterial pneumolysin. Although there is very little sequence similarity between these proteins – there is only one amino acid conserved throughout all the known bacterial and vertebrate proteins of this family, a glycine at a critical position for the conformational change – the structures are similar enough to suggest that the proteins all once had a common ancestor.

Saibil and her colleagues used electron microscopy to discover that this protein forms a pore through a similar mechanism to pneumolysin: the helical region that unfolds into the beta hairpin to form the pore forms the core of the molecular machine and is largely unchanged between the structures. There are some differences between the structures, however; in particular, there is no need for the perforin structure to ‘collapse’ as the molecule has ‘arms’ that are long enough to form the hairpin and punch the hole without bending into an arch.

The mechanism through which the NK cells kill their target cells is now quite well understood. When the two cells come into contact they form a temporary structure called an immune synapse that allows the pore to form and proteases called granzymes, which induce apoptosis, to enter the target cells. This YouTube video illustrates the natural killer cells’ mechanism of action, and this one shows a detailed view of the immune synapse. Other, similar proteins have been identified in oyster mushrooms; these form more rigid structures that are easier to work with. Saibil’s group and their collaborators have been able to solve the structure of this protein in intermediate stages of pore formation and are beginning to gain an understanding of exactly how it unfolds.

The pore of the oyster mushroom protein pleurotolysin, a member of the pneumolysin family. Image © Natalya Lukoyanova and Helen Saibil, from Lukoyanova et al., PLoS Biology 13:e1002049

Mutations in perforin that prevent it from functioning cause a rare disease called haemophagocytic lymphohistiocytosis, which is almost invariably fatal in childhood. Understanding the mechanism of action of this important family of protein ‘weapons’ in both attack and defence may help find a cure for this devastating condition, as well as for some commoner disorders of the immune system and important infectious diseases.

This post is cross-posted from the Birkbeck Events blog.

PPS students can learn much more about electron microscopy by taking the second-year module Techniques in Structural Molecular Biology to complete the MSc.