Our lab is fascinated by the question of how small molecules like proteins, lipids and nucleotides self-assemble into cells and tissues that are thousands and millions of times large than molecular dimensions. How do the molecules know where they are, and whether the structures that they have made are the right size and shape? By combining highly sensitive techniques to visualize and manipulate individual biological molecules, with theory and modeling, we are trying to understand the interaction rules that allow molecules to work together to form highly organized and dynamic cellular structures.
Extensive Research Description
How do cells measure length and time? How do they count? How do they sense their shapet? These are some of the most important outstanding questions in cell and developmental biology. For example, how do proliferating cells know how big they are and when to stop growing and to start dividing? How do organs, such as brain and liver, ensure that they manufacture enough cells, but not too many? How do cells control the dimensions of their organelles, such as endosomes, cilia and mitotic spindles? How do cells make certain that they have exactly two copies of each chromosome, that they have just one cilium, but that dividing cells have exactly two mitotic-spindle poles?
These are difficult questions because they address “systems-level” processes whose length-scales and time-scales differ strikingly from those of the constituent molecules—the proteins, lipids and nucleic acids—that form the molecular building blocks of cells. The goal of our research is to find molecule-level answers to these systems-level questions. Just as a radio or a computer must be understood in terms of their constituent analog or digital circuits, cell and tissue morphology must be understood in terms of underlying molecular circuits. These circuits comprise networks of interacting molecules. What kinds of computations can they perform? How do they assemble? What feedback-control mechanisms operate to control the size, shape and organization of molecular assemblies? Reaction-diffusion equations have dominated our thinking about biological patterning since the seminar work of Turing in the 1950s; however, we argue that active, mechanical processes mediated by motor proteins and the cytoskeleton, may play an even more important role in morphogenesis than purely chemical mechanisms (Howard et al. 2011).
Our research is focused on the mechanics of the cytoskeleton, an internal filamentous scaffold that provides structure to the cell by supporting the plasma membrane and anchoring the organelles. The cytoskeleton serves as a network of tracks along which motor proteins transport subcellular structures. The research is concentrated on the mechanics of the cytoskeleton (Howard, Mechanics of Motor Proteins and the Cytoskeleton, Sinauer and Associates, Sunderland MA, 2001), with a particular emphasis on microtubules and microtubule-based motor proteins.
On the one hand, we are interested in how these proteins work as nanomachines. How do motors and other microtubule-binding proteins couple an energy source, the hydrolysis of ATP or GTP, into mechanical work used to regulate the assembly and disassembly of the cytoskeleton? On the other hand, we are interested in how microtubules and their motors move and shape organelles and cells. For example, how do the dynamic properties of microtubules drive the assembly of the mitotic spindle and control its size and shape? And how does dynein drive the beating of the axoneme, leading to the serpentine motion of sperm and cilia?
Our approach is to combine measurement and theory. The challenge is that the complexity of biological systems makes quantitative measurements and their interpretation exceedingly difficult. The key to circumventing these difficulties is the use of single-molecule techniques, in whose development our lab played an important role (e.g. Howard et al. 1989). Using single-molecule optical techniques, the interactions between the individual motor and cytoskeletal molecules can be characterized in vitro and in vivo. These interactions constitute a form of mechanical signaling (Howard 2009). We then use theory, primarily from statistical physics and non-linear dynamics, to predict how the individual interactions lead to the collective behavior of ensembles of molecules. We then test these predictions with quantitative in vivo experiments on intact cells.
Microtubules are biological polymers that alternate between periods of growth and shrinkage. This process, termed dynamic instability by its discovers Mitchison and Kirshner, is crucial for microtubule length regulation, for the exploration of intracellular space, and for cellular force generation. Despite its central role in cell biology, dynamic instability is poorly understood. For example, the GTP-cap, which is thought to regulate catastrophe, the transition from growth to shrinkage, has never been visualized on dynamic microtubules.
We are advancing the field by bringing three new approaches to the problem. First, we have developed assays to study microtubule dynamics using single-molecule techniques: total-internal reflection fluorescence microscopy (Brouhard et al. 2008, and see right for motors moving along a microtubule) and optical tweezers (Bormuth et al. 2009, Jannasch et al. 2013, Trushko et al. 2013). Second, we have used these assays to figure out how microtubule-associated proteins regulate microtubule dynamics. We have shown how depolymerizing kinesins target microtubule ends and couple ATP hydrolysis to microtubule shortening (Helenius et al. 2006, Varga et a. 2006, 2009, Friel and Howard 2011); we discovered that XMAP215 is a processive polymerase (Brouhard et al. 2008, Widlund et al. 2011); and we found that the end-binding protein EB1 recognizes the nucleotide state of tubulin (Zanic et al. 2009) to increase catastrophe and to synergize with XMAP215 to increase microtubule growth rates (Zanic et al. 2013). The third approach is to use theory to gain insight into microtubule length control (Varga et al. 2009), the catastrophe switch (Gardner et al. 2011, Bowne-Anderson et al. 2013) and the collective properties of motor proteins (Leduc et al. 2012).
The motility of cilia and flagella
That the motor protein dynein drives microtubule sliding and bending in cilia and flagella has been known since the classic experiments of Gibbons, Brokaw and Satir in the 1960s and 1970s. However, how the key question of how the activity of the dyneins is coordinated to give a periodic beat remains an open. We are bringing a new constellation of techniques to the problem. In earlier work, we hypothesized that the dyneins are coordinated through a force-sensing mechanism: sliding forces provide positive feedback that switches the activity of the motors across the axis of the axoneme (the motile structure within cilia and flagella) and couples the motors along the length of the axoneme (Riedel-Kruse et al. 2007, Howard 2009). Using high-speed imaging and high-precision image analysis, we showed that our hypothesis can account very satisfactorily for the beat of mammalian sperm.
We are now using the single-celled alga Chlamydomonas Reinhardtii as a model system to study (i) the beating of isolated, reactivated axonemes, and (ii) the activity of individual dynein molecules purified from these axonemes. In this way we hope to obtain a molecular understanding of one of the most elegant and enigmatic examples of cell locomotion. Oscillators are a paradigms of emergence because the system behavior obscures the contributions of the individual players. We hope to the obtain the first truly molecular and biochemical understanding of a cellular oscillator.
We are working on several other projects in which the dynamic and structural properties of microtubules underly important cellular processes such as mitosis (Grill et al. 2003, Pecreaux et al. 2006, Redemann et al. 2010, Reber et al. 2013), mechanotransduction (Howard and Bechstedt et al. 2004, Bechstedt et al. 2010, Liang et al. 2013) and neuronal morphology.
- Howard J (2014) Quantitative cell biology: the essential role of theory. Mol Biol Cell. 25:3438-3440. doi: 10.1091/mbc.E14-02-0715. PMID: 25368416
- Geyer VF, Jülicher F, Howard J, & Friedrich BM (2013). Cell-body rocking is a dominant mechanism for flagellar synchronization in a swimming alga. Proc. Natl. Acad. Sci. USA doi:10.1073/pnas.1300895110.
- Zanic Z, Widlund PO, Hyman AA, Howard J. (2013) Synergy between XMAP215 and EB1 increases microtubule growth rates to physiological levels. Nat. Cell Biol. 15:688-693 (DOI: 10.1038/ncb2744)
- Bowne-Anderson H, Zanic M, Kauer M, Howard J. (2013) Microtubule dynamic instability: A new model with coupled GTP hydrolysis and multistep catastrophe. Bioessays 35: 452-61. PMID: 23532586
- Liang X, Madrid J, Gärtner R, Verbavatz JM, Schiklenk C, Wilsch-Bräuninger M, Bogdanova A, Stenger F, Voigt A, Howard J. (2013) A NOMPC-Dependent Membrane-Microtubule Connector Is a Candidate for the Gating Spring in Fly Mechanoreceptors. Curr. Biol. 23:
- Leduc, C., Padberg-Gehle, K., Varga, V., Helbing, D., Diez, S. & Howard, J. (2012) Molecular crowding creates traffic jams of kinesin motors on microtubules. Proc. Natl. Acad. Sci. U S A. 109: 6100-6105.
- Gardner, M.K., Zanic, M., Gell, C., Bormuth, V., Howard, J. (2011) Depolymerizing kinesins Kip3 and MCAK shape cellular microtubule architecture by differential control of catastrophe. Cell 147: 1092–1103.