Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

The overall goal of this florescent time-lapse microscopy protocol is to provide a method for studying biological cell processes that underpin the development and cellular differentiation in the sporulating filamentous bacterium, Streptomyces venezuelae. The describe method provides an excellent platform to study cell biological processes that are central to Streptomyces lifecycle, including dynamic protein localization, polarized growth and sporulation septation. The main advantage of this technique is that Streptomyces venezuelae sporulates in liquid, which allows us to grow the cells in a microfluidic device and microscopically monitor the complete lifecycle.

This method demonstrates the immense potential of Streptomyces venezuelae as a new developmental system for the genus, because it allows for live cell imaging of the differentiation of a multicenter mycelium into chains of spores. To begin, inoculate 30 milliliters of supplemented growth medium with 10 microliters of spores from the S.venezuela strain to be imaged. For consistent growth and sporulation of the cells, use a baffled flask or a flask that contains a spring to allow for sufficient aeration.

Culture the cells for 35 to 40 hours at 30 degrees Celsius and 250 RPM. When ready, you should be able to see the mycelial fragments and spores via liquid-mounted phase contrast microscopy. Centrifuge one milliliter of the culture in a tabletop centrifuge at 400 times G for one minute to pellet the mycelium and the larger cell fragments.

Then transfer approximately 300 microliters of the supernatant containing a suspension of spores, to a new 1.5 millimeter tube and placed the tube on ice. Retain the remaining culture medium for use in a later step. Next, dilute the spores one to 20 in supplemented growth medium, and keep the diluted spores on ice until they are needed.

Pour the remaining culture medium in a 50-milliliter beaker and then draw up 10 milliliters with a syringe and use a sterile 22 micrometer syringe filter to filter sterilize the remaining culture medium. To obtain spent supplemented growth medium that is free of spores and mycelial fragments. Keep the filtered spent supplemented growth medium for a few days at four degrees celsius if additional experiments, using similar growth conditions, are to be conducted.

Each microfluidic plate can be used for up to four independent experiments. To avoid contaminating the unused flow chambers, make sure you use sterile solutions and working conditions when setting up the experiment. Begin by removing the shipping solution from the microfluidic plate.

Then rinse the wells with sterile supplemented growth medium. Once rinsed, add 300 microliters of the supplemented growth medium to the inlet of well one, and 300 microliters of the spent supplemented growth medium to the wells two through six. Next, load 40 microliters of the diluted spores to well eight of lane A and seal the manifold to the plate in accordance with the manufacturer's instructions.

Launch the microfluidic's control software and select the appropriate plate type. Set up a flow program to flow medium from inlet wells one through five at six psi for two minutes per well in order to prime the flow channel and the culture chamber. Then have it flow supplemented growth medium at six psi into inlet well one for six hours.

This will allow germination and vegetative growth to take place. Have the program switch after six hours to the spent supplemented growth medium and flow it into wells two through five at six psi for the remainder of the experiment. Pre-warm the environmental chamber to 30 degrees Celsius in advance.

Then turn on the microscope and the microscope control software. Put a high numerical aperture oil immersion objective into place and confirm that the appropriate filters and diagrammatic mirrors set to acquire differential interference contrast images, along with the images of the yellow florescent and the red fluorescent protein fusions are in place. Place a drop of immersion oil into the objective and to the bottom of the imaging window on the microfluidic plate as well.

Carefully mount the sealed microfluidic device onto the stage of the inverted microscope and secure it into place. Use the embedded position markers to bring the imaging window of the microfluidic culture chamber into focus. Focus on the leftmost part of the first flow chamber labeled A, and then move the stage to trap size five, corresponding to the trap height of 7 micrometers.

In the microfluidic software, set the system to load cells from inlet well eight at four psi for 15 seconds. After the process is run, check the cell density in the culture chamber by moving the stage across the imaging window. If no spores were trapped, repeat the cell loading step or alternatively, increase the loading pressure and/or time until the desired cell density of one to 10 spores per imaging window is achieved.

Take care to avoid overloading the culture chamber. Next, start the previously prepared flow program in the control software and allow the microfluidic plate to heat equilibrate for one hour in the microscope stage before starting image acquisition. In the microscope control software, set up a multidimensional acquisition to take multiple images at multiple stage positions over time by first specifying a directory for the automatic saving of the image files.

Next, go to the illumination settings and enter predetermined optimal illumination settings for each specific construct. Then set up a time series to acquire images every 40 minutes for 24 hours. In order to determine stage positions and set the autofocus, scan the culture chamber and store stage positions for each imaging position of interest.

Ensure that the single-stage positions are located far enough apart to minimize photo bleaching and photo toxicity. Once the Z coordinates of the selected stage positions are verified, activate the hardware autofocus. Then start the time-lapse experiment in the microscope control software.

Stop the image acquisition after 24 to 30 hours, or when the hyphae in the region of interest have differentiated into spores. Then stop the flow program in the software and disassemble the microfluidic device. Prepare the used microfluidic plate for short-term storage by removing any remaining medium from the inlet wells, the waste well, and the cell loading well.

Then fill up the used wells of lane A and wells of the unused lanes with sterile PBS. Finally, seal the plate with pear film to prevent it from drying out. And store the plate at four degrees Celsius.

The successful live cell imaging of the entire S.venezuela lifecycle yields a continuous time series, including the key developmental stages of germination, vegetative growth and sporulation. During germination or vegetative growth, DivIVA-mCherry exclusively accumulates at the growing hypnal tips or marks newly forming hyphal branching points. In contrast, FtsZ-YPet forms single ringlike structures at irregular intervals in the growing mycelium.

These structures provide the scaffold for the synthesis of nonconstructive vegetative cross walls leading to the formation of interconnected hyphal compartments. In sporulating hyphae, the localization pattern of FtsZ-YPet changes dramatically. First, helical FtsZ-YPet filaments tumble along the hypha, and then in a sudden, almost synchronous event, these helices coalesce into a ladder of regularly spaced FtsZ-YPet rings.

Finally, sporulation septa becomes discernible in the differential interference contrast images and eventually, the new spores are released. This technique provides a robust protocol to perform live cell imaging of the complete Streptomyces lifecycle. The microfluidic system is easy to use.

It offers experimental flexibility and it allows long-term monitoring of the This experimental set up also offers a great starting point to investigate specific developmental events in response to changing cultural conditions or the use of florescent dyes to monitor peptidoglycan synthesis or to visualize chromosome organization.