Microscopy Facility

Learn more about the microscopy facility at Central Michigan University and how you can start researching with cutting edge equipment right on campus.

If you have questions about our microscopy facility, reach out to one of our experienced personnel members.

NameDepartmentOfficePhone NumberEmail
Philip OshelBiologyBiosciences 1304989-774-3576philip.oshel@cmich.edu
Joanne M. DannenhofferBiologyBiosciences 3408989-774-2509 danne1jm@cmich.edu
Philip L. HertzlerBiologyBiosciences 3406989-774-2393philip.l.hertzler@cmich.edu
Jonathan KeltyBiologyBiosciences 4103989-774-1382kelty1jd@cmich.edu
Eric W. LintonBiologyBiosciences 3401989-774-3969eric.linton@cmich.edu
Anna MonfilsBiologyBiosciences 2401989-774-2492monfi1ak@cmich.edu
Daniel E. WujekBiologyProfessor EmeritusN/Adaniel.e.wujek@cmich.edu

Equipment

Nikon A1R CLSM

The Nikon is a confocal microscope, meaning it images only the in-focus light coming from the sample, the out of focus light is rejected.

It uses samples that are fluorescent by labeling target structures (like mitochondria) or molecules (like specific proteins) with a fluorescent molecule. The fluorescing targets then are only seen where they exist in the cells. The samples are then scanned by lasers of a specific wavelength, so only those fluorescent targets that are excited by that wavelength of light are seen.

Also, the genome of a cell can be modified by added DNA that encodes a fluorescent protein, like Green Fluorescent Protein or Red Fluorescent Protein. This DNA is added to the beginning or end of the DNA that makes a specific protein, so that the GFP is added to the target protein. This means the fluorescence only shows up when the target protein is made.

Since only in-focus light is accepted by the imaging detectors, and only fluorescing molecules are seen, the use can then take images at different depths within a sample - the Z-dimension in an X-Y-Z graph - and the images can be stacked (this is called a Z-stack) and combined to create a 3D reconstruction of the volume of the sample. Like an entire nucleus, with its contained proteins and DNA/RNA, instead of just 2D slices of the nucleus.

Olympus Fluoview 300 CLSM

Students use our Olympus Fluoview 300 CLSM ​​every day to learn confocal microscopy.

Our Confocal Laser Scanning Microscope​ is based on an Olympus BX50 upright microscope, and has Differential Interference Contrast optics and two extra-long working distance water immersion objectives on a special two position turret. A microinjection system for electrophysiological studies is also available.

This microscope is used in the Bio 553 Confocal Microscopy class. It uses three PMT detectors, two for epi-confocal imaging, and one for transmitted-light imaging. Imaging capabilities include reflected-light confocal and transmitted-laser DIC imaging. Argon 488 nm and He/Ne lasers are installed on the microscope.

Objective Lenses

4X, 10X, 20X, 60X Plan Apochromat DIC; 40X Plan Fluorite DIC; 20X Plan Fluorite Long Working Distance Phase; 20X and 40X Plan Fluorite Water-immersion DIC

Micromanipulators

Narishige hydraulic hanging joystick-controlled fine and coarse micromanipulators

Hitachi 3400N-II Scanning Electron Microscope

Scanning Electron Microscope are basic research instruments widely used across academia and industry to do everything from research to creating art.

SEM's have the widest variety of uses of any microscope.

This scanning electron microscope (SEM) is the newest addition to the Microscopy Facility. The Hitachi 3400N-II has a tungsten filament electron gun with a large, motorized specimen chamber and has variable-pressure capability and an IR chamberscope. It is also equipped with a Deben cooling stage. There are several detectors on this instrument: the usual secondary detector, a 5-element backscattered electron detector, an environmental secondary electron detector, a Thermo-Noran System Seven energy-dispersive X-ray spectrometer with a silicon-drift detector, and a Gatan color cathodoluminescence detector.

The biology department offers an O Scanning Electron Microscopy Techniques class on a yearly basis during the spring semester. This is a laboratory-intensive class, with its focus on a final project, chosen by each student, which includes theory on SEM imaging and operation.

Procedures

Preparatory Equipment

Samples for viewing in microscopes, particularly biological samples, usually require special preparation methods. This is especially true of samples to be viewed in electron microscopes, where they must withstand high vacuums and bombardment with high-voltage electron beams.

These sample preparation methods typically use specialized instruments, available in the Imaging Facility:

  • ​​Ladd Research vacuum evaporator for carbon or metal evaporation.
  • Denton Desk II sputter coater with either gold or 60/40 gold/palladium target.
  • Polaron "bomb" critical-point dryer.
  • 1 RMC Powertome and 2 MT-2B Sorvall ultramicrotomes.
  • RMC GMK glass knife maker.
  • Standard chemicals and equipment for fixing and embedding samples for confocal microscopy, SEM, and TEM.
  • Equipment for freeze-fixation by plunging into liquid nitrogen and for making and plunge-freezing into slush nitrogen.

Please contact Philip Oshel by email or by phone at 989-774-3576 for information or consultation on use of the preparation equipment, specimen preparation, and design of studies using microscopy.

Light Microscope Cleaning Procedures

Microscopes are sophisticated instruments that require periodic maintenance and cleaning. Dust, lint, pollen, dirt, and oil can cause a deterioration of image quality.

​A few guidelines to follow:

  • NEVER mix immersion oil types.
  • ALWAYS clean immersion oil off oil lenses when you are finished with a scope (or at the end of a lab or end of the day).
  • Immersion oil that is left on a lens or mixed with other oils can solidify and ruin the lens.
  • Do not use cotton swabs (Q-tips and the like) to clean lenses! The cotton fibers will scratch lens coatings. Dacron and foam-tipped swabs are OK.

Major Component Routine Cleaning

Oculars
  1. Routine disinfection of ocular to prevent transmission of conjunctivitis. Using a piece of non-linting lens paper soaked in 70% ethanol, lightly blot the surface to disinfect. Do not saturate the ocular! Air dry or use bunched lens paper to soak up the excess.
  2. Routine cleaning of oils and fingerprints and other contamination. First, use a duster and gently blow off the lens. Then, start with a bunched-up piece of non-linting lens paper dipped in lens cleaner (purple solution, aka Sparkle) and lightly blot the lens. Soak up any of the excess solution on the lens surface with fresh paper.
  3. Do not let the lens cleaner air dry! Remove it with a fresh, non-linting lens paper, very gently wiping the remaining lens cleaner from the lens, moving the paper in one direction (left to right, up to down, or the like). Use as much lens paper as needed to do the job.
Objectives

Clean following the same procedure above in Oculars. Extreme care should be given to cleaning the objectives, and this should only be done by an experienced user. If you are unsure about cleaning objectives, ask Philip Oshel in 024c Brooks Hall.

Stage
  1. Clean first with 70% ethanol and a Kim-wipe (not paper towels).
  2. If residue or contamination still remains, use the lens cleaner on a Kim-wipe to further clean the stage.
  3. Care must be taken not to saturate the moving parts of the stage when cleaning!
Body
  1. In general, the body of the microscope shouldn't need to be cleaned.
  2. The scopes are cleaned completely once a year.
  3. If it needs cleaning right away, follow the guidelines for cleaning the stage.

If the image does not improve or if you are unsure about any of the above procedures, please notify Philip Oshel in 024C Brooks Hall or at (989) 774-3576.