This post was contributed by Dr Clare Sansom, senior associate lecturer in the Department of Biological Sciences. Dr Sansom attended Dr Anthony Roberts’s lecture during Birkbeck Science Week 2016.

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.

Walking molecules and their importance for human health

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.

Snapshots

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.

A single leg with two feet

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.

The structure of Dynein

Dynein

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.

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.

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This post was contributed by Dr Clare Sansom, senior associate lecturer in the Department of Biological Sciences. Dr Sansom attended Dr Maya Topf’s lecture on Computational Modelling during Birkbeck Science Week 2016.

Brain computer (copyright Marcos Fernandez via Flickr. Image cropped)

The Wednesday of Birkbeck Science Week – 13 April – was set aside to celebrate women in science, and it included a talk by Maya Topf of the Department of Biological Sciences. Maya, who was educated in Israel and Oxford, came to Birkbeck on an MRC fellowship after a post-doc at UCSF and has rapidly worked up the academic ladder to the position of reader in computational biology. She will be appointed as a full Professor in October this year.

Maya began by explaining that her research involves making models: specifically, three-dimensional models of biological molecules. Models have enabled scientists to make sense of biological processes since Watson and Crick’s double helical model of DNA showed how this molecule could both replicate itself and act as a template for the synthesis of proteins. This model, celebrated in the film Life Story that was shown earlier in Science Week, would not have been possible without the X-ray photographs of DNA fibres obtained by Rosalind Franklin, then working at King’s College.

The purpose of computational modelling

And the main purpose of modelling molecules is the same now as it was in the 1950s: to discover how they function, and specifically how they function in the environment of the cell. We still have no means of observing what protein molecules – the tiny ‘machines’ that drive all cellular processes – look like when they are at work; all we have is models that may be more or less precise. The very first protein structures to be determined were of the oxygen-carrying proteins myoglobin and haemoglobin, and the first of these, published in 1960, were very imprecise: it was possible to see the shape of the chain but no individual atom positions. These, and all early protein structures, were obtained by X-ray crystallography; ten years later the same group used the same technique to determine a structure in which all atoms except hydrogens could be seen.

These two proteins have now also been studied using two other structural biology techniques, nuclear magnetic resonance and, most recently, electron microscopy. This last technique is best suited for studying large proteins and complexes of many protein chains, and therefore not suitable for studying most forms of haemoglobin, a small, simple protein. Haemoglobin in earthworms, however, functions as a complex of many individual molecules. Electron microscopy gave a low-resolution picture of the overall shape of these molecules, much like those first haemoglobin structures, and a more precise picture was built up by ‘docking’ atomic-resolution X-ray structures of a single haemoglobin molecule into the shape of the fold.

During the last half-century these three techniques have generated structures for a wide range of proteins, leading to insights in many areas of biochemistry: how the body’s catalysts, the enzymes, work; how drugs bind to their receptors; and how a ‘large’ molecular complex, the ribosome, can synthesise all the proteins that a cell needs from RNA templates. The first atomic structures of this ‘molecular machine’ were obtained in the early 2000s and have transformed our view of protein translation since then (see these videos from the Howard Hughes Medical Institute in the US: basic and more advanced versions).

But, as real proteins are too small to be visible with even the best light microscopes, we need to realise that even these experimental structures are models. Each of the three techniques has its own advantages and limitations. X-ray crystallography needs protein crystals, which can be difficult or even impossible to obtain for particular proteins; electron microscopy cannot be used to study small proteins, but NMR works best with these. All three techniques are complex, time-consuming and expensive, and therefore proteins with known structures are greatly outnumbered by those without structures. There are probably about 43,000 known structures of ‘distinctly different’ proteins known compared to over half a million well-characterised protein sequences.

Bridging the sequence-structure gap

Maya explained that much of her group’s work concerns trying to bridge this ‘sequence-structure gap’ by using computers to model unknown protein structures. There are several ways of doing this; if the computers are powerful enough and the molecule is small enough (and the smallest proteins can be) it is possible to generate a model structure ‘from first principles’ using physics. These techniques assume that the molecules are likely to occupy conformations in which their energy is low. The best results simulate protein folding to produce model structures that can be very close to the experimentally-determined ones, but these require an enormous amount of computational power. Less expensive computer modelling methods tend to rely more on experimental data; Maya collaborates with Helen Saibil in Biological Sciences to fit atomic structures of individual proteins to lower-resolution maps of protein complexes that were generated by electron microscopy. Proteins studied in this way include GroEL, a ‘molecular chaperone’ that forms a chamber that isolates unstructured proteins so that they can fold.

Dr Maya Topf

Another method of modelling protein structures uses evolution, and relies on the fact that there are remarkably few different basic protein structures – each of the 43,000 known protein structures takes up one of only about 1,000 different folds. Just as all birds have the same basic pattern, with two legs and two wings, all proteins with a particular function will usually have a similar fold. It is therefore possible to model the structure of a protein based on one or more of its evolutionary relatives, in a technique called ‘homology modelling’. In some cases, it is possible to produce a usable model from the structure of a related protein from a very different type of organism. It was more than a decade after the publication of the first bacterial ribosome structures before similar structures could be obtained from mammalian ribosomes, but many useful results were obtained during that time by modelling mammalian ribosome sequences using the bacterial structures and low-resolution electron microscopy data.

Maya ended her talk by stressing that structural biology is a science of model-building. It requires experimental data complemented by physics and by evolution, and, almost above all, it requires powerful computers. Generally, the more sources of information can be combined into a model, the nearer the ‘correct’ structure that model will be: and to quote the statistician George Box, ‘all models are wrong, but some are useful’.

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