Kate Luby-Phelps, PhD, director of the Live Cell Imaging Facility at the University of Texas Southwestern Medical Center, talks to contributing editor Tanuja Koppal, PhD, about some of the different techniques available for live cell imaging and the improvements in recent years when it comes to hardware and software tools. She also highlights some of the common challenges encountered during live cell imaging and ways to ensure that the cellular conditions during imaging remain close to physiological conditions.
Q: Can you tell us about your core facility, the instruments you have, and the work you do?
A: Our live cell imaging core, established in 2004, is a fee-for-service facility, and we serve the entire UT Southwestern Medical School campus. We typically train our 200 or so users, and they use our facility for an hourly fee. We have four laser scanning confocal microscopes, and three of them are equipped for multiphoton imaging. We also have two spinning disk confocal microscopes, several wide-field fluorescence microscopes, and a superresolution instrument configured for 3-D structured illumination microscopy and 2-D localization microscopy. We also manage two image-processing workstations with specialized software for image deconvolution and for 3-D image analysis.
Q: What types of training do you provide to your users?
A: Our training is one-on-one and very basic in terms of teaching them what they need to know. If it’s something new and complicated, then we work together to make sure all the kinks are worked out. We have a number of image-analysis software tools that we provide for complicated analysis, but what we usually recommend for basic analysis is free downloadable software like ImageJ and Fiji. We try to make image analysis accessible to everybody; we do this through basic training, and I also teach workshops and courses. We do give advice on sample preparation, but until recently we have not done any sample prep ourselves. Our users bring their samples, which are frozen or paraffin-cut sections or tissue culture cells, and for immunofluorescence we sometimes help them with fixation and staining. For live cell imaging, people come with their cells and we advise them on how to plate the cells, typically using 35 mm glass-bottom dishes, as they work better for optics with oil immersion lenses. Very rarely we have people working with cells plated in glass-bottom microtiter plates, as those tend to be very expensive.
Q: What improvements have you seen in recent years when it comes to hardware, software, and reagents for live cell imaging?
A: The biggest change has been the development of new technologies for imaging, which has made it possible to image many more types of samples with better resolution. The development of light sheet fluorescence microscopy has made it possible to look through large transparent samples like zebrafish. It allows you to focus in on parts of the animal or organism without worrying about stray light coming in from other parts of the sample. The development of super-resolution technologies has also been a big game-changer. The increased accessibility of techniques like multiphoton microscopy, which can be used to look at brains of live animals like mice, is also a big help to the average user.
Q: How do you ensure that the cellular conditions during imaging remain close to physiological conditions?
A: For live cell imaging, I think it’s important that the cells or animals are happy on the microscope stage and do what they would normally do in the incubator or cage. For cells, organoids, and organs, that would mean keeping them at the right temperature, pH, and humidity. For live cell imaging, the microscopes are enclosed in an incubator. It’s a plexiglass chamber that encloses the stage and the optics and is maintained at specific physiological conditions. With samples like cultured organoids, finding these conditions to keep the cells happy can be very tricky. At our core, we encounter just about any type of tissue culture cells for imaging. Cells, if they start blebbing or changing shape, then they are not happy. Sometimes the cells may look good, but they could be getting light damage from the strong illumination used. My criterion is, if they can go through two rounds of cell division then they are probably healthy. We have been able to image the same batch of stem cells for two weeks and get cell lineage data. We also have a microscope that is equipped to do live animal imaging in rodents or rabbits. It’s mostly mice or rats that we image, or sometimes we do get organs in culture. However, the people in our facility do not have the expertise to make sure that the animals under anesthesia are doing fine, so our users have to take care of that themselves. The main thing is to keep the animal warm and adequately sedated.
In live cell imaging, focal drift can also be a big issue. It may seem like a small improvement, but for time-lapse imaging, the technology for maintaining focus over long periods of time has been very important. While everything has become a little more complicated in imaging, it has become a lot easier to find out when there is a problem as there is an obvious change or error message reported. Microscopes that are automated have checks and balances in place to make sure that the hardware and software are communicating with each other properly. It is easy to troubleshoot when there is a minor problem, but for other things we have service contracts in place. A lot of the vendors now offer remote sessions, where they can guide us step-by-step to try to figure out what the problem might be and how it can be fixed.
Q: What advice can you give users working on live cell imaging?
A: My advice to people who are looking to buy a new imaging instrument is to have the vendor bring the instrument to your lab and demonstrate how it works using your samples, in your environment. Second, don’t forget to get the hardware for autofocus, especially if you are doing time-lapse experiments, because it makes a huge difference. For live cell imaging you are also going to need some kind of incubation chamber. There are two types of incubators—one that sits on the microscope stage and the other that sits around the stage and includes the lenses. One big challenge is that the lens, which includes a big chunk of metal, sucks all the heat away. So if you don’t heat the lens, you have a temperature difference between the sample and the lens, particularly with an oil immersion lens where you have direct contact between the lens and the glass on which the sample is kept. To avoid that, you will need a lens heater along with a stage-top incubator or you will need an incubator that encloses the lens as well. Maintaining the right focus and the right temperature are two of the most important things in live cell imaging.
Another piece of advice is to use a # 1.5 (0.17 mm) coverslip if you are using an oil immersion lens for highresolution imaging, because all the optics are optimized for that thickness. You will not get the best results if you don’t use a coverslip with that particular thickness. The other challenge is the trade-off between the working distance and the resolution. Working distance is the distance between the front lens of your objective and the sample. For a thick sample you really need a confocal microscope, and you need a lens with a big working distance. In terms of equipment that I would like to buy, there is a super-resolution microscope that has much better resolution than other techniques. It is called a stimulated emission depletion microscope, and it can cost more than a million dollars!
Kate Phelps has a PhD in cell biology, with over 30 years of experience in microscopic imaging and more than 50 publications involving light and electron microscopy. She was trained in electron microscopy as an undergraduate at Yale University and she did her doctoral work at the University of Colorado. Her postdoctoral training was in the laboratory of Dr. D. Lansing Taylor, a pioneer in the application of modern fluorescence microscopy to biomedical research, including live cell imaging. Kate has been director of the University of Texas Southwestern Live Cell Imaging Facility since 2004. Since 2012, she thas also been the director of the UT Southwestern Electron Microscopy Facility.