The institute provides laboratories and 24 specialised suites for more than 200 scientists and research students. Learn more about our facilities and equipment below.
Microscopes are used to produce magnified visual or photographic images of small objects. Samples can be easily visualised without labelling e.g. brightfield, phase contrast. Further characteristics can be determined via fluorescence labelling.
In epi-fluorescence/widefield microscopy, the whole specimen is bathed in light and all parts of the specimen are excited at the same time. Therefore, out of focus light originating from above and below the focal plane (secondary fluorescence) is captured along with light originating from it.
Confocal microscopy uses a screen with a pinhole to exclude out of focus light and produce a sharper image. This can generate an image through a point scanning process, where it scans each pixel individually and compiles a digital image. It can also be used to obtain images of planes at various depths within the sample (known as optical sectioning or z stacks) and then create a 3D image. In spinning disk confocal there are numerous pinholes which are moved by means of a rotating disk. Therefore, the entire image is collected at the same time. This increases speed, but sacrifices resolution compared to point scanning confocal microscopes.
Spectral unmixing can be used to eliminate autofluorescence and distinguish overlapping signals when multiple fluorophores are used. This process often increases sensitivity and accuracy in multi-parametric immunofluorescence.
Some epi-fluorescence and confocal microscopy applications include:-
- Localisation of proteins
- Live cell imaging of cellular processes such as endocytosis, phagocytosis, protein trafficking
- Cell-cell interactions
- Complex techniques for measuring cell complexity (fluorescence recovery after photobleaching; FRAP), or to determine molecular proximity (fluorescent resonance energy transfer, FRET).
- 3D reconstruction
We conduct regular training sessions on all our microscopy equipment (see below).
For further details please contact
Connie Jackaman: email@example.com
Flow cytometry is a technique for counting and examining cells or particles (resolution: approx. 0.2 μm to 100 μm). It allows simultaneous multi-parametric analysis of thousands of particles per second. When a sample in solution is injected into a flow cytometer, the particles are randomly distributed and the sample must be ordered into a stream of single particles to be interrogated by the machine’s detection system. This is via either hydrodynamic focussing or acoustic focussing.
Hydrodynamic focussing of cells into single file occurs by pressure differential between sheath fluid and sample. Acoustic Focussing uses ultrasonic waves (over 2 MHz, similar to those used in medical imaging) generated by a piezoelectric device to position cells into a single, focused line along the central axis of a capillary.
Benchtop analysers are used to interrogate cell populations, however cannot collect cells or particles post-analysis. High-speed cell sorter flow cytometers are used for physically sorting cells or particles of interest for further assays.
Some flow cytometry applications include:
- Identification and quantification of different cell types by surface and/or intracellular proteins
- Cell cycle analysis
- Analysis of phosphorylated proteins
- Cytokine measurement via cytokine bead arrays
- Mitochondrial membrane potential
- Bacterial and marine biology analysis
We conduct regular training sessions on all our flow cytometry analysers (see below).
For further details please contact
Jeanne Le Masurier: firstname.lastname@example.org
- Attune acoustic flow cytometer
- BD FACS Canto II flow cytometer
- BD LSR Fortessa flow cytometer
Cell sorting is provided as a service to users on the BD FACS Jazz (see below).
Please contact core facility staff for further details and for any cell sorting requests.
- BD FACS Jazz
Brochures to download
For sample processing the facility provides tissue culture and histology facilities. Tissue culture is used to grow cells by simulating conditions (in vitro within the lab) that may occur within the body. Cells can be cultured as an immortal cell line or directly from tissues or blood. Histological techniques provide a visual means for the examination and analysis of cell/tissue physiology. This can include embedding and sectioning tissues so that they can be visualised. Samples processed using tissue culture or histology are often analysed within the flow cytometry, metabolic analysis and microscopy facilities.
Tissue culture facility
There are 4 levels of tissue culture activity mainly based on Mycoplasma status. Mycoplasma is a bacteria which can significantly impact on cell growth and interpretation of biological results.
- “Quarantine” work i.e. primary human cultures which have not been tested for any known human pathogens (most human work), viral work, parasite work and Mycoplasma status positive cell lines which are being treated.
- “Dirty” cell lines which are Mycoplasma status unknown, only one Mycoplasma negative result or irregularly tested for Mycoplasma and primary cultures which are negative for known human pathogens (most animal-based work from the SPF facility).
- “Clean” cell lines which have been regularly tested for Mycoplasma and have two negative results minimum.
- “Ultra Clean” cell lines which have been regularly tested for Mycoplasma and have three negative results minimum and cultured without antibiotics.
We conduct regular induction sessions for tissue culture.
For further details please contact the tissue culture core facility staff member.
- Leica processing and embedding station
- for paraffin blocks
- Dewaxing and staining workstation
- Microme cryostat
We conduct regular training sessions on the above histology equipment.
A cell’s ability to utilise energy efficiently is critical for a number of cellular processes. For analysis of bioenergetics, the Seahorse Extracellular Flux Analyser can be used to investigate the two major energy producing pathways in cells – oxidative metabolism and glycolytic metabolism in live cells. Cells can be examined for oxygen consumption rate (OCR), and extracellular acidification rate (ECAR), in order to assess cellular functions such as oxidative phosphorylation, glycolysis, and fatty acid oxidation.
Some applications for live cells include:
- Simultaneously measurement of OCR and ECAR
- Rapidly detect cellular responses to stimuli, substrates, inhibitors, and other drugs.
- Stress tests to assess mitochondrial function
Further details are below and for further details please contact the tissue culture core facility staff member.
- Seahorse Bioanalyser
All instruments have high-end software which can be used for analysis. However, priority of these instruments is for data collection rather than analysis – if they are not being using for data collection they can be booked for analysis.
- NIS elements software (with object tracking) on the Nikon A1+ confocal microscope
- Volocity 3D analysis software on the Ultraview Vox confocal microscope
- FACS Diva on the FACS Canto II and Fortessa flow cytometers
- Attune cytometric software on the attune flow cytometer
An offline analysis workstation is available (see below). NIS elements software are also available as a sign in/sign out dongles (contact Connie Jackaman: Connie.Jackaman@curtin.edu.au for use).
- Networked computer for data collection point
- Microscopy image analysis
- Nikon Advanced Research software with deconvolution
- Image J software
- Flow cytometry data analysis
- FlowJo analysis software
- FCAP Array software
- Data presentation
- Microsoft W7
- GraphPad Prism
We conduct regular training sessions on microscopy and flow cytometry data analysis.