MLS Microscopes

Different Contrast Generation Techniques

SMBS Microscopes

Common use microscopes
Researcher-specific microscopes

Microscope maintenance

Promoting “microscope” awareness

Different Contrast Generation Techniques

Many techniques, eg:

• Phase contrast
  • requires specific condenser and objectives
• DIC (Differential interference contrast)
  • requires specific condensers, prisms etc
• Hoffman-Modulation Contrast
  • requires specific condenser and objectives
  • good for plastics

All require the illumination system to be aligned.

• Colour staining of specimen
  • requires transmitted white light source (Halogen lamp).
• Fluorescence staining of specimen
  • requires epi-fluorescence light source (Mercury lamp), excitation (Ex) and emission filters (Em).

Microscopes

Common use Microscopes within the School

• Rm 1.14 MLS Bldg
• Upright
• B&W CCD camera attached
• V++ image acquisition software
• Used for: epiflourescence mainly
  • Does have Normarski capability also
• Four fluorescence channels:
  • A: GFP, Alexa 488, Cy2
  • B: Cy3, Rhodamine, TR
  • C: DAPI
  • D: Cy5
• Objectives: 10x, 20x, 40xoil, 60xoil, 100xoil, 100xoil

• Located in Rm 1.14
• Upright microscope
• Fujix Colour camera, CCD cooled camera attached
• Photograb image acquisition software used
• Used for: Colour imaging with Normarski. eg. ISH, H&E
• Tissue prep: Slides optimal
• Objectives: 10x, 20x, 40x, 60x,100x oil
• Normarski filters can be moved between objectives
  

• Rm 1.14
• Stereo Microscope with Fluorescence filters: GFP, CFP, YFP
• Olympus DP70 camera attached
• AnalySIS Life Science Research imaging acquisition software
• Uses? Viewing and dissecting GFP-positive tissue
  • Not sufficient resolution for viewing live cells best suited for embryo tissue

3rd Floor Microscope Room, MLS Building

Nikon TE 300 (RHS)

• Rm 3.03
• Inverted microscope
• CoolsnapFX B&W CCD Cooled Camera attached
• V++ imaging acquisition software
• Flourescence filters: FITC, TR/Cy3, DAPI
• Objectives:
  • 4x, 10x, 20x, 40x (silver)- optimal for Flourescence;
  • 20x and 40x (black)- optimal for Hoffman Modulated Contrast (HM)
• Uses?
  
  • HM optimal for imaging through plastic (i.e. live cells)

  • Other objectives used for imaging fixed and stained cells and tissues in TC dish

Nikon TE300 (LHS) 

• Rm 3.03
• Inverted microscope
• CC12 Soft Imaging System Camera
• AnalySIS Life Sciences Research Image Acquisition Software
• Fluorescence filters: FITC, TR/Cy3, DAPI
• Objectives: 4x, 10x, 20x, 40x
• Uses? Colour imaging through plastic (e.g. whole-mount ISH, live cells), fluorescence; Normarski optics

4th Floor, MLS Building

BioRad MRC 600 (4th Floor Confocal)

• Rm 4.03
• Inverted microscope
• Lasers: Kr/Ar
• Objectives: 10x, 40x, 60x, 100x
• 2 channel
• FITC/Texas Red Fluorescence imaging of thick tissue, 50 µM or more.
  • Relatively low resolution, may not be suitable for publication quality images

Researcher-specific microscopes

Managed by the research lab that owns the microscope

Heavy use by those labs requires that they have priority useage

Specialised setups

All have CCD cameras

Various combinations:

• Epi-fluorescence
• DIC
• Hoffman
• Incubators
  • Temp
  • CO2
• Z drive

Various Software packages:

• Timelapse
• FRET
• Deconvolution

• Zeiss Axiophot, located in Rm 1.14 (Decon)
• State-of-the-Art microscope with up to 8 Fluorescent filters, Computer controlled microscope + imaging
• Ideal for thick tissue specimens
• objectives: 2.5x, 10x, 20x, 63xoil, 100xoil

• Olympus IX81, located in Rm 2.41
• Inverted microscope with Deconvolution capacity
• Full range of pbjectives and fluorescent filters

• Olympus IX70, located on Level 4
• DAPI, FITC, TxRed
• Phase, 4X, 10X, 20X, 40X oil, 100X oil

• Level 3, Leica Stereo Microscope with DFC 480 camera attached

Maintenance of microscopes within the School

Olympus AX70

Chris Cursaro, Technical Officer, Adelaide Proteomics, Rm 1.51, Ext: 37521

Nicholas Eyre, Postdoctoral Researcher, Michael Beard's Lab, Micro& Immuno, Rm 4.16, Ext: 37551

Zeiss Axiophot (Screen attached)

Chris Cursaro, Technical Officer, Adelaide Proteomics, Rm 1.51, Ext: 37521

Tanya Henshall, PhD Student, Richards Lab, Rm 2.14, Ext: 35752

Edwina Sutton, NH&MRC Postdoctoral Research Officer, Paul Thomas' Lab, Rm 3.09, Ext: 35009

Morgan Newman, PhD Student, Lardelli Lab, Rm 1.24, Ext: 34863

Zeiss Axioplan2 (Decon)

Stephen Gregory, Genetics and CMGD Research Officer, Saint Lab, Rm 1.32, Ext: 37536

3rd Floor Microscope Room, MLS Building

Nikon TE 300 (RHS)

David Bersten, PhD Student, Whitelaw Lab, Rm 3.08, Ext: 34806

Fook Hing, PhD Student, Jensen Lab, Rm 3.42, Ext: 33791  

Nikon TE 300 (LHS)

James Hughes, PhD Student, Rathjen Lab, Rm 3.34, Ext: 34623

Contact person(s) for each microscope...  

• Provide a point of contact for training and trouble-shooting minor problems
• Are able to identify and then defer major problems to Chris Cursaro
• Organise microscope “checklist” to be put up at the microscope
• Ensure that there are adequate consumables i.e. lens tissues etc at each microscope and that the area is generally clean
• Monitor data levels and organise data removal if necessary
• Replace Log books when necessary

Promoting “microscope” awareness

Rule number 1:

Learn to use the Microscope you need to use well before you need to use it.

Rule Number 2:

If in any doubt, STOP, get expert help !

Rule Number 3:

Clean up equipment appropriately and remove your mess.