Facilities and Techniques
Our lab primarily utilizes confocal microscopy using an inverted Nikon TE2000 epifluorescence widefield microscope. We employ this microscope through the use of the in vitro motility assay, an assay in which we are able to image fluorescently labeled actin filaments sliding over a bed of myosin molecules in a flow cell. This set up allows us to manipulate a wide variety of experimental conditions and perturbations to probe the mechanics of both unregulated actin-myosin interactions and thin filament-myosin interactions. Our primary image capture device is a Roper Cascade 512B CCD camera. Our microscope is also equipped with a 488/561 laser which we can use to perform TIRF microscopy, a form of microscopy where an evanescent wave is generated that selectively illuminates the surface of a coverslip which effectively blocks the background fluorescence generated by the flow cell under normal widefield conditions. Using these tools we have the resolving power to image as small as a quantum dot (nanometer spatial resolution) up to whole cell dimensions. Classically these techniques are used to measure the collective behavior of muscle molecules acting in ensemble, however their use is not limited to this alone.
In addition to typical microscopy and TIRF, we have at our disposal several other techniques to augment our ability to examine molecular events. Using a class IV 1064nm UV laser, we're able to use light to manipulate and trap small molecules which allows us to measure force generation at the level of a single molecule by observing the displacement generated by a single myosin head against a load (Laser Trapping). We also have at our disposal a Stanford Photonics XR/Turbo-G camera that can resolve events on a near-millisecond timescale (1000 frames/sec), allowing us to visualize a host of events in real time such as the subsecond regulation of muscle that are not typically observable.
While microscopy is a large focus of the Baker lab, it is only one of many techniques employed by us. Another, through the collaboration of Christine Cremo's laboratory, is stopped flow. Stopped flow is a rapid mixing device used to study the chemical kinetics of a reaction in solution. Currently, it is one of the most important and frequently used techniques in the field regarding kinetics research. Two or more solutions containing reagents of interest are rapidly driven into a high efficient mixer, mixed, and are then studied by any experimental methods suitable. Just prior to stopping the flow a steady state is achieved. What is amazing is that the reaction entering the flow cell occurs and can be resolved on the millisecond timescale. This allows us to measure the kinetics of a fast reaction. The stop syringe limits the volume injected, hence the term “stopped flow”. In our lab, we specifically use stopped flow to measure the kinetics of pyrene labeled actin (both regulated and unregulated) binding to a single headed (S1) myosin under varying conditions.
Two more techniques employed by our lab are the use of ATPase assays and spindown centrifugation binding assays. ATPase assays, while not specific to a muscle laboratory, are an essential tool. This assay allows you to measure the rate of ATP turnover, and in the four state model of actin-myosin binding, myosin binding is essentially an ATPase reaction. The reason we measure this reaction is because Ton, the time myosin spends strongly bound to actin, is a function of both ADP release and ATP binding. This parameter is critical because in a detachment limited model Ton is the determinant of both muscle velocity and force generation at a given step size.
In our sedimentation assays, we analyze how various proteins including myosin will bind to F- (filamentous) or G- (monomeric) actin. Using centrifugation, proteins that bind to the filament form of actin will co-sediment and form a pellet at the bottom of the centrifugation tube, while proteins that don't bind actin or only bind the monomeric form will remain in the supernatant. This technique can further be extended to analyze the binding affinity of a protein to actin - a vital property when considering the weak and strong binding properties of myosin and the binding of the regulatory troponin and tropomyosin to actin.
As well as being able to utilize a variety of experimental techniques, we also utilize a variety of muscle proteins. Our lab is proficient in the isolation and purification of all three types of muscle: cardiac, skeletal, and smooth. While our animal model is typically a rabbit, we've isolated proteins from tissue as small as a mouse's heart to as large as a bovine's. In addition to the muscle proteins myosin and actin, we're also proficient in the purification of many of the other proteins present in the muscle sarcomere, including but not limited to troponin, tropomyosin, myosin light chain kinase, calmodulin, and calponin. This wide array of proteins and animal models further bolsters our arsenal of experimental conditions available to experiment on.
Finally, while a great deal of our time is spent at the bench doing experiments, we also employ a large amount of our resources developing computer simulations of our theoretical models to test hypotheses and behaviors we observe in our experimental data. In addition, we work on developing and refining current and innovative analysis techniques to describe the behavior we see in our experiments using a wide variety of mathematical and statistical methods.