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HeLa cell in mitosis.

Bioimaging

Light microscopy based imaging from the single cell through to the whole organism.

The Bioimaging Core Facility provides access, training and support for state-of-the-art AFM, widefield, confocal and high-content live cell imaging.

We provide access to a wide range of modern light microscope systems from Atomic Force Microscopy (AFM) through to in vivo multi-photon confocals.

This means we have a microscope for almost every imaging application and are happy to discuss your requirements and arrange training/access on the most appropriate systems.

In addition to The University of Manchester’s researchers, we also support external access.

 

On this page:

 

Neurones labelled with the 'Brainbow' of fluorescent proteins in a mouse brain.
 

How we work

We give support throughout your imaging project from experiment design through to image acquisition, processing and data analysis.

Microscopy projects

We encourage new users to discuss their microscopy projects with us before initiating work.

This allows us to advise on experiment design, sample preparation and to discuss the various imaging options.

Once we have agreed on the best system/s to use we provide full and ongoing user training. This ensures that all our users can use the systems independently and to their full potential.

Slidescanning service

We also offer a slidescanning service where the slides are automatically and fully digitised in either colour brightfield or up to four fluorescent channels.

 

Applications

If you have an imaging project, it is likely that we have the microscope you need.

We have a wide range of microscope systems that offer advanced imaging capabilities.

If you have an imaging project, even if it’s new and novel, then it is likely that we have at least one microscope that can meet your requirements. If it isn’t available, we’ll work with you to make it available.

Typical experiments include:

  • Live cell imaging of cells over 1 to 72 hours using widefield, spinning disc or point scanning confocals to allow tracking and protein expression studies.
  • Colocalisation and expression patterns of fixed cells and tissue.
  • Protein-protein interactions investigated using FCS, FLIM and super resolution microscopy.
  • Protein dynamics studied using photokinetic events such as FRAP, FLIP and photoactivation.
  • Structural characterisation and mechanical sensing using biosensors (FLIPPER and FRET/FLIM).
  • Ultra-structural properties of cell and tissue using Atomic Force Microscopy (AFM).
Green fluorescent protein in the growing root tip of Arabidopsis thaliana.
 

Technologies and equipment

A cutting-edge imaging project requires access to state-of-the-art imaging equipment.

Constant investment in the Bioimaging Core Facility ensure that our equipment remains cutting-edge and allows our users access to the latest imaging techniques and technologies.

The facility is spread over three purpose-built suites in the Michael Smith and Stopford Buildings. We have five full-time members of staff who provide full training and support on a wide range of imaging techniques.

  • Atomic force microscopy (AFM)
  • Point scanning confocals (super resolution/FLIM/FCS/in vivo MP)
  • Spinning disc and lightsheet confocals
  • High content live cell imaging
  • Widefield (snapshot, deconvolution, TIRF, laser microdissection, bioluminescence, protein patterning, plate reader)
  • Digital slides scanning

Find out more about the equipment available for each category.

 

Atomic force microscopy (AFM)

Atomic Force Microscopy (AFM) involves the raster scanning of a cantilever-bound probe over a surface and measures the deflection of the cantilever at each position to produce a 3D image of the sample with near atomic resolution.

In addition to imaging the AFM can also be used to perform force spectroscopy. Here the probe is moved in the Z direction towards the sample surface only, indenting slightly on contact before retracting again. The deflection of the cantilever is used to calculate the force applied to the probe at each displacement, along with adhesion forces as the cantilever retracts. In this way the forces involved in polymer folding/unfolding, biomolecular interactions and the compression of biomaterials (including cells) can be measured.

Point scanning confocals (super resolution/FLIM/FCS/in vivo MP)

Confocal microscopy works by rastering a single diffraction limited laser spot across the sample so that excitation and detection is achieved “point by point” to build up the complete image. The use a confocal pinhole prevents out of focus light from reaching the detector and provides a complete 3D volume of the sample that is free from blur with a high level of resolution.

Super Resolution is achieved using Stimulated Emission Depletion (STED) microscopy. A scanning laser is used to excite the fluorophore as in a regular confocal system, but a second “doughnut” shaped laser is superimposed with this laser and causes a depletion of fluorescence inside that area. This STED effect takes the resolution below the diffraction limit for that wavelength and provides super-resolution images.

FCS/FLIM are techniques that allow the confocal system to measure protein interactions and dynamics by following changes in their mobility within the image beam (FCS) or the time the protein remains in the excited fluorescent state (FLIM).

Two-photon confocal microscopy requires two photons of long wavelength excitation light to arrive simultaneously at the fluorophore to cause excitation. The only place with a density of photons high enough for this to occur is at the focal plane and as a result there is no need for a confocal pinhole to reject out of focus light as there isn’t any out of focal plane excitation. This non-reliance on a pinhole to achieve optical sectioning and use of long wavelength excitation light makes the two-photon microscope ideally suited for deep tissue imaging and in vivo experiments.

Spinning disc and lightsheet confocals

Spinning disk confocals use an array of micro-lenses to split the excitation laser light into many smaller beamlets that scan simultaneously across the sample. The emitted light then passes through a matched spinning disc of pinholes that reject the out of focus light and produce a confocal image. Since this technique uses hundreds of beamlets, rather than a single rastered laser, production of confocal images and z-stacks is therefore faster and less likely to cause photo-bleaching or cell damage than normal confocal, making spinning discs the ideal choice for confocal imaging of live cells.

Light Sheet microscopes uses an orthogonal arrangement of camera and illumination to create an 3D confocal stack of the samples. This is achieved using a thin sheet of laser light (1um to 14um thick) to illuminate just the focal plane of the objective, thus there is no need for a confocal pinhole as just fluorophores within the focal plane are illuminated. By moving the sample quickly through this optical arrangement of plane illumination and imaging, it is possible to achieve high speed confocal volumes with very low photobleaching.

High content live cell imaging

These systems are designed for maintaining cells for long periods of time by maintaining a stable environment of temperature, humidity, and CO2. Motorised XYZ stages, laser autofocus, and intelligent software allows multi-dimensional experiments to be carried out easily on multi-well plates and advanced analysis software is able to batch process these data sets to provide unbiased results on growth patterns, drug treatments and transfection studies. A range of imaging techniques are available from label free phase imaging, widefield fluorescence and AiryScan confocal imaging on whole plates over several days of imaging.

Widefield (snapshot, deconvolution, TIRF, laser microdissection, bioluminescence, protein patterning, plate reader)

Snapshot

  • 3 x Zeiss AxioImager Z1 fluorescence microscopes
  • Leica M205 stereo fluorescence microscope
  • Olympus BX63 with colour camera

Although these are the simplest microscope systems that we have, they are also our brightest and take great pictures so it may be that you don’t need to use the more complicated microscopes to answer your scientific questions. The microscopes are manual and very easy to use.

 

High end

Laser microdissection: uses a focused UV laser to cut around cells and small areas of tissue mounted on membrane slides. This cut tissue is then picked up using special sticky-cap Eppendorf tubes. Thus small regions of the samples can be cut and separated from the rest of the tissue without any physical handling of the sample and is suitable for downstream applications in the Genomic Technologies and Mass Spectroscopy core facilities.

Deconvolution: takes a series of widefield images through the sample, each of which contains both the in-focus light and out-of-focus information for each z-position.  A mathematical algorithm is then performed on this data set to relocate the out of focus light back into its correct focal plane to generate sharp, bright, free from blur optical stack. However, since none of the light was rejected during the imaging process, the technique is more sensitive than regular confocal imaging and is ideal for samples where the fluorescence is weak.

Primo protein micro-patterning: allows the user to create any pattern and/or gradient of proteins on a coverslip based dish. These can then be seeded with cells and imaged on any of our microscope systems.

TIRF: Total Internal Reflection (TIRF) microscopyis a technique that allows the imaging of fluorophores very close to the coverslip while the rest of the sample remains “dark” producing high contrast images with very low photo-toxicity. The angle of the incident laser is adjusted to cause total internal reflection at the coverslip-sample buffer boundary. While this ensures that the sample is no longer excited by a penetrative laser beam, an evanescent illumination field is generated close to the coverslip on the sample side. This evanescent field excites any fluorophores within only a few 100nm of the coverslip while the remaining volume remains unexcited and thus permits imaging of proteins in contact (or very close proximity) to the coverslip.

Fluorescence plate reader: This system does not produce images but instead gives a fluorescent read out for each well. It can be used with 96 or 384 well plates to measure values for the following: Fluorescence Absorbance (200 to 1000nm), Fluorescence Emission Intensity (250nm to 850nm), Fluorescence Polarization, Time-Resolved Fluorescence and Luminescence. The system is temperature controlled and also has an 8 channel fluidics module for the automatic addition of drugs or reagents during the time-course.

 

Bioluminescence

Bioluminescence does not require an excitation light source in order for the protein of interest to glow. Instead, bioluminescent or chemiluminescent signals are produced within the cells/tissue. These can be imaged in live cells over very long time periods (hours-weeks). However, the weak signal emitted from these samples requires very long exposures and the system needs to be light tight to prevent ambient light from entering the system. The system has a 35mm dish holder/heater an environmental control chamber (for 37°C and 5% CO2) all contained in a light tight enclosure.

Digital slidescanning

We have two 3D Histech Pannoramic P250 which can automatically detect and digitise up to 250 slides in a single run.

The slides can be brightfield or 4-channel fluorescence and images can be analysed using 3D Histech free slideviewer software or free software such as QuPath.

Our Olympus BX63 with scanning stage can be used to create high resolution images where the user is able to defined both the region of interest and its focus map.

This is then scanned and stitched into a single large image which can be opened in most free software such as QuPath and Fiji ImageJ.

 

Publications and outputs

High impact publications often rely on access to state-of-the art imaging systems. The Bioimaging Core Facility provides those systems.

We are an essential resource that contributes to numerous high impact publications every year.

Here are some of the recent highlights.

 

Endosomal recycling tubule scission and integrin recycling involve the membrane curvature-supporting protein LITAF.

Journal of Cell Science, 2021.

A high impact paper showcasing the range of microscopes available in the facility, from simple snapshot through to confocal microscopy and live cell imaging.

The paper also shows the power of combining light and EM microscopy.

Quantitative single-cell live imaging links HES5 dynamics with cell-state and fate in murine neurogenesis.

Nature Communications, 2019.

Understanding how a complex system is co-ordinated and controlled requires a knowledge of the number of molecules involved.

This paper uses FCS to accurately predict this number of molecules and then image cells over several days to follow their fate.

Glucocorticoids rapidly inhibit cell migration through a novel, non-transcriptional HDAC6 pathway.

Journal of Cell Science, 2020.

FCS, live cell imaging and modelling showcase the techniques available in the facility and how they can be combined to produce a high impact publication.

Relief of talin autoinhibition triggers a force-independent association with vinculin.

Journal of Cell Biology, 2019.

Combining live cell imaging and photokinetic capabilities of the systems to understand the mechanisms involved in cell adhesion and migration.

Phase contrast image of live tissue cultures cells growing on extra-cellular matrix.
 

External access

Although primarily focused on supporting research at The University of Manchester, we are also able to support work from other academic institutions and industry.

Academic institutions

We are happy to provide access to all our microscope systems and slidescanning service to users from any academic institution.

We will provide these users with the same level of training and support as we would for our internal users.

Contact us for further details.

Industry

We have worked for a number of multinational companies, both on an individual basis and as part of packages of work involving multiple facilities.

Work may be on a collaborative basis or purely as a service provision.

All work is fully documented and contractual, and non-disclosure compliance is assured.

Please contact our Business Development Manager for further details if you are interested in accessing the facility

Dr Joanne Flannelly
Email: joanne.flannelly@manchester.ac.uk

The localisation of 9 different proteins in a developing Drosophila melanogaster embryo.
 

Contact us

Get in touch if you require any further information, or would like to inquire about using our facility.

Dr Peter March, Principal Technologist

Email: peter.march@manchester.ac.uk
Telephone: +44 (0)161 275 1571

Bioimaging Core Facility
The Michael Smith Building
The University of Manchester
Oxford Road
Manchester
M13 9PT

Maps and travel

We are based in the AV Hill Building (Building 71 on the University campus map).

See: University maps and travel section

Technology platforms

We have a pioneering environment and facilities for research, innovation and technology development.

Technology platforms main page

Cross facility support