Author Archives: Anthony Hayes

IN FOCUS: Virtual Microscopy Image Database: An Update.

You may remember one of our blogs from 2015 about a virtual histology slidebox  in development by the Bioimaging Research Hub? If not, link here.  Well, I’m very pleased to report that we’ve made considerable progress since then.

The resource has now been moved from its humble beginnings (a Rasberry Pi/raid drive set-up) to a new PC server based within the Bioimaging Research Hub. The database has also been developed significantly through mySQL which allows efficient management of the image metadata via a web interface, allowing the images to be sorted, filtered and navigated online.  

The image collection has now been expanded significantly; thus, in addition to the original histopathology collection (which contained approximately 400 digitised sections of normal and pathological tissues), we now have two new additional sections on cell biology and parasitology.

The cell biology section contains both zoomable/navigable images and interactive 3D models of intracellular structure, including major organelles, cytoplasmic inclusions and cytoskeletal components. These were all generated from confocal fluorescence datasets imaged using our Zeiss LSM880 airyscan confocal system and rendered in 3D via Bitplane Imaris. The parasitology section, meanwhile, contains over 200 new zoomable/navigable  images of various parasitic species,  organised phylogenetically for easy reference and sorting. As before, these images were generated using our Navigator slide scanning system.

So far, the database has been trialed for small group anatomy teaching, as well as for a number of BIOSI practical classes including Research Techniques (#BIT002), Advanced Research Techniques (#BI4002) and the Identifying Organelles (#BI2231) module. It is also utilized extensively for outreach and engagement activities within the Bioimaging hub to showcase our research capabilities.

As the database is a bespoke system that has been developed in house, there are no costly subscriptions involved. We are also uniquely positioned within the Bioimaging Hub to expand and develop the database according to the needs of the user.  It therefore has enormous potential as a centralized repository for microscopical image data for teaching, research and outreach/engagement purposes.

We are planning to add additional sections on plant biology and entomology  and we would welcome collaboration with any BIOSI staff who have access to the relevant slide resources and would be happy to  help in curating these collections.

Ultimately, the plan is to find a permanent home for the virtual microscopy database as part of the new e-learning and assessment facility (eLEAF) within BIOSI.

Please take a look at the database here: http://vmdb.bios.cf.ac.uk/. Any feedback (+/-) would be welcome.

Thanks again to all involved so far.

AJH

IN FOCUS: 3D Pollen Prints Not To Be Sniffed At: Printing Pollen for the Met Office.

Above:  Not to be sneezed at: 3D pollen prints for the Met Office (grass, green; oak, yellow and birch, blue).

Disclaimer: If you suffer from hayfever then please avoid spending too long on this page – it may be detrimental to your health!

I bet you didn’t know that one in five people  suffer from hayfever and that 95% of pollen sufferers are allergic to grass pollen in the UK alone? Well neither did I until I visited the Met Office’s pollen forecast website. 

It seems that some of the worst offenders are pollen grains from grass, oak and birch which play havoc with the mucous membranes during the pollen season, causing sneezing, nasal congestion, itchy eyes and triggering asthma in susceptible individuals (and to make matters worse,  these conditions are exacerbated  by drinking alcohol – so no respite there!)

Having read some of our previous blogs (here and here), the Met Office recently asked the Bioimaging Hub if we would generate 3D printed models of some of the worse culprits  (shown above) for their outreach & engagement programme to help promote awareness of hayfever. 

The 3D prints were generated from surface-rendered confocal microscope volume datasets with help from BIOSI 3D printing. We’ve used the technique to generate physical models of a variety of microscopic samples ranging in size from subcellular organelles to whole developmental organisms. If you’re interested, then further details of the methodology are available below.

AJH

Further reading:

IN FOCUS: Getting to the Root of the Problem.

Above: ‘Crowning glory’: Webcam shots (1-12) showing stages in the process of 3D printing a giant human molar (left) and the resultant 3D print with support scaffold removed (central and right).

The other day we were presented with a problem: was it possible to generate a 3D model of a human tooth that could be used for dental teaching and outreach purposes? The only thing was, the individual concerned didn’t specify the desired size. With a build volume of 215 x 215 x 300mm and printing resolution of 20-200 microns, our new Ultimaker 3 Extended 3D printer can print BIG, so what better application to put the new instrument through its paces! After a quick search on Thingiverse.com, we downloaded a stereolithography (.stl) file of a human molar tooth segmented from computed tomography (thanks to fvillena). We decided to print it as big as we could, but using the lowest print resolution and lowest level of infill. The results, shown above, are quite impressive – it took approximately 24 hours to print the tooth (crown-side down, root-side up) and with the support scaffold removed resulted in a 3D model approximately 300mm in height – about the same size as tooth from an adult T-Rex!! I suppose we can now be accused of (wait for it…) getting a bit long in the tooth!

AJH

Further reading:

NEWS: Microscope Maintenance Course: Keeping your ‘Scope in Tip-Top Condition.

Above: Some of the class microscopes in various states of dismantlement.

It goes without saying, to get the very best out of a microscope you need to know how to optimise and maintain its performance. That said,  you’d be surprised just how many microscopists don’t know how to properly set up and maintain their microscopes.

Recently, we run our first  microscope maintenance course as part of Cardiff University’s Continuous Professional Development (CPD) programme.  We can’t tell you who it was for; but suffice to say, they use microscopes a lot in their work. The two day course covered the basics of light microscopy and the procedures necessary to keep a microscope squeaky- clean and correctly aligned. The practical element of the course saw delegates clean, rebuild and align both upright widefield and stereo-zoom microscopes. 

Pleasingly, the course was well-received with very good to excellent feedback. Thanks to all who participated on the two busy but  enjoyable days. Thanks must also go to our undergrad students for soiling and mis-aligning the microscopes ahead of the course – they did a far better job than we ever could ; )

Further reading:

AJH.

SPOKE EQUIPMENT: New Ultimaker 3 ‘Extended’ 3D Printer.

Above: The new Ultimaker ‘Extended’ 3D printer building a stage insert for a novel microscope system.

The Bioimaging Research Hub has recently purchased an Ultimaker 3 ‘Extended’ professional 3D printer. The printer will allow users to 3D print bespoke pieces of scientific equipment or generate scale models  of microscopic samples for use in their research as well as for teaching, outreach and engagement activities (examples can be  found  here and here).

The 3D printer utilises FDM (Fused Deposition Modelling) printing technology and has a range of advanced features allowing the fabrication of professional quality, high resolution 3D prints. The printer can print in two different colours or a single colour in addition to a dissolvable PVA support scaffold, thus allowing complex overhanging structures to be printed at high fidelity whilst significantly reducing  finishing time.

The printer has a large build volume (215 x 215 x 300mm), supports a range of materials (nylon, PLA, ABS, CPE and PVA) and has a print resolution of 20-200 microns.  The printer is wi-fi enabled with an internal webcam so that users can remotely monitor the progress of their 3D prints.

Further details of the equipment are available through our equipment database.

Further reading:

AJH

CORE EQUIPMENT: New Zeiss Lightsheet Z.1 Microscope.

Above: The new Zeiss Lightsheet Z.1 fluorescence microscope.

Above: Tripping the light fantastic: the new Zeiss Lightsheet Z.1 fluorescence microscope.

A state-of-the-art, Zeiss Lightsheet Z.1 system has recently been installed within the Bioimaging Research Hub (BIOSI 2; E/0.03). The single plane illumination microscope (SPIM) , purchased through generous funding by Cardiff University’s Research Infrastructure Fund (Lead applicant: Dr Pete Watson), allows fast and gentle multi-channel 3D/4D fluorescence imaging at the sub-cellular level. The system offers the potential of whole organ imaging of fixed cleared samples and can provide an unprecedented insight into developmental processes occurring in live model organisms such as Arabadopsis thaliana (Thale Cress), Danio rerio (Zebrafish) and Drosophila melanogaster (Fruit fly), and in vitro organoid at a high spatio-temporal resolution. Further information about this system is available via the Bioimaging Hub equipment database.

Webinars:

Further reading:

SPOKE EQUIPMENT: Zeiss PALM MicroBeam Laser Micro-dissector.

palm

Above: ‘Cutting-edge’ equipment: the Zeiss PALM MicroBeam laser micro-dissector located at the European Cancer Stem Cell Research Institute (ECSCRI)

A Zeiss PALM MicroBeam laser micro-dissector system is now available as a spoke of the Bioimaging Research Hub. The equipment, which is located at the European Cancer Stem Cell Research Institute (ECSCRI),  allows isolation of DNA, RNA and protein from laser micro-dissected samples from both histological sections (paraffin wax or cryo) and live cells. Further details of the system are available through our equipment database. All enquiries for this system should be directed towards Mr Mark Bishop.

AJH

CORE EQUIPMENT: New Spinning Disc Confocal System for Live Cell Imaging.

spinning-disc

Above: The new spinning disc confocal system

The old electron microscopy darkroom (BIOSI 2; E/0.05) within the Bioimaging Research Hub has recently been refurbished as a live cell imaging suite via generous support from BIOSI. It now houses a spinning disc confocal system for  fast, live cell imaging applications. The system is based around an Olympus IX71 inverted microscope, kindly provided by Dr Pete Watson, which has been upgraded, via ISSF funding, with a Crest Optics X-Light V2 confocal head, a Cairn Research tri-line laser bank (405nm, 488nm, 561nm) and a Hamamatsu ORCA Flash 4 sCMOS digital camera with M-View Gemini image splitter.  The system is fully integrated via Molecular Devices MetaMorph software and boasts a 40″ 4k display.  The system will expand the Hub’s imaging toolbox, enabling high speed, multi-position, multi-colour 3D/4D image acquisition. Support systems for live cell imaging (i.e. gas and incubation) are also available within the facility. Further details of this system are available through our equipment database.

AJH

IN FOCUS: Cutting Through the Fog: Reducing Background Autofluorescence in Microscopy.

Autofluorescent bone sample

Above: Autofluorescence from mixed connective tissues imaged by confocal microscopy (left). The autofluorescent emissions can be spectrally-resolved through wavelength scanning (right). Excitation at 488nm.

Whilst autofluorescence from endogenous fluorophores can reveal much about the biochemical composition of a sample, it can also hamper the microscopic detection of targeted fluorochromes if they emit light at the same wavelengths as endogenous fluors. Indeed, without proper controls, complex background autofluorescence can lead to misinterpretation of image data and generation of false positive results.

Autofluorescence derives from multiple sources within the sample – the main culprits are  NADH and NADPH, lipofuscins, flavins, elastin and collagen (and lignin and chlorophyll in plants). The excitation and emission ranges of the worst offenders have been shown below. It follows that tissues with high collagen and elastin contents, e.g. skin, tendon and cartilage, autofluoresce very brightly; as do tissues that are rich in metabolic breakdown products such as lipofuscin, e.g. liver, spleen etc.

Autofluorescent data

Adding to the problem is the effect of chemical fixatives (e.g. formalin, glutaraldehyde etc) and solvents used to preserve tissue architecture for microscopy: the cross-linkages generated by these chemicals increase autofluorescence, which can be worsened further by long-term storage of the fixed processed tissues.

So, dear reader, here’s some simple advice on steps that you can take to address this common problem:

1. Include an unlabelled control to evaluate the level of autofluorescence within your sample.

  • Observation of unlabelled samples through RGB fluorescent filters (note their transmission characteristics) will help identify where in the visible spectrum the autofluorescent signal is brightest.
  • Spectral (lambda, wavelength) scanning will allow you to precisely identify the fluorescent emission spectra from endogenous fluorochromes and can help separate their emissions from those of your fluorochrome (see above figure).

2. Select fluorochromes that are outside the range of the autofluorescence.

  • If the autofluorescence signal is high in the blue, then move into the green; if it’s high in the green, move into the red – or better still, the far red (if your system can detect in this range).
  • Use modern fluorescent probes (e.g. Alexa Fluor, Dylight, or Atto range) instead of first generation fluorochromes.  They are brighter, more photo-stable and have narrower excitation and emission bands. They are also available in variants that span the near UV, visible and far red range of the spectrum, affording you plenty of choice.

3. Use a microscope with filters optimised for your choice of fluorochromes.

  • Band-pass filters which collect emissions within a specific range may be more useful than long-pass filter sets which collect all emissions past a certain wavelength. The narrower the range of the band-pass filter, then the better it can separate fluorophores with close emission spectra.

4. If the autofluorescence is unevenly distributed within your sample, use targeted microscopy to avoid it.

5. If you can’t avoid the autofluorescence, then take measures to remove or reduce it.

  • Analyse the pixel intensity distribution within your image and try thresholding out the lower intensity autofluorescence signal.
  • Pre-bleach your samples in a light box using a high intensity illumination source prior to fluorescent labelling (see below reference)
  • Treat samples with a chemical reagent (e.g. sodium borohydride, Sudan black B, ammonium ethanol etc) to reduce background autofluorescence (see below reference)

6. If all else fails, consider the following:

  • use cryoprocessed material as an alternative to chemical fixation and paraffin wax processing.
  • avoid long term storage of material/archival tissue samples.
  • try a different detection modality (e.g. immunoperoxidase instead of immunofluorescence)

AJH

Further reading

Wright Cell Imaging Facility. Autofluorescence: Causes and Cures

 

IN-FOCUS: Better To Burn Bright Than To Fade Away: Reducing Photo-bleaching in Fluorescence Microscopy.

[Parameter-Settings] FileVersion = 2000 Date/Time = 0000:00:00 00:00:00 Date/Time + ms = 0000:00:00,00:00:00:000 User Name = TCS User Width = 1032 Length = 1032 Bits per Sample = 8 Used Bits per Sample = 8 Samples per Pixel = 3 ScanMode = xy Series Name = demo2.lei

Above: Photo-bleaching (fading) occurs when a fluorochrome permanently loses the ability to fluoresce due to photon-induced chemical damage and covalent modification. 

Hands up if you’ve spent hours preparing a sample for fluorescence microscopy only to see the signal disappear before your eyes upon excitation? Frustrating eh (unless, of course, FRAP is your objective)? Well here’s some simple and sound advice on how you can minimise photo-bleaching and get the best out of your samples under the fluorescence microscope.

1. Visualise your samples immediately after fluorescent labelling – this is when they are at their brightest.

  • If this is not possible then loosely wrap your samples in aluminium foil and keep them in the dark at 4oC until you get the opportunity to image them.

2. Minimise their exposure to light in order to reduce photo-bleaching.

  • visualise your samples under low light conditions.
  • use transmitted light to find a region of interest (ROI) and then switch to epifluorescence observation – avoid dwelling too long on the ROI.
  • step down the intensity level of excitation light or insert a neutral density filter into the light path.
  • set up imaging parameters on a neighbouring region and then return to the ROI for image capture.
  • use image binning to reduce exposure time.
  • use the microscope shutter to switch off the light source between images.
  • create a photo-bleach curve from a timed series of images. This can be used to normalise for loss of fluorescence intensity.

3. Switch to a mounting medium with anti-fade protection e.g. Vectashield, Prolong Gold/Diamond, SlowFade Gold/Diamond. These work by reducing the oxygen available for photo-oxidation reactions, thus reducing photo-bleaching. N.B. Many of these are available with a nuclear counterstain (e.g. Dapi) included in the formulation. Alternatively, make your own anti-fade reagent (instructions below).

4. Switch to brighter, more photo-stable fluorochromes. First generation fluorochromes such as FITC and TRITC photo-bleach readily (and are pH sensitive) thus should be replaced with modern dyes such as the Alexa Fluor, Dylight, or Atto  range of fluorochromes, which are much brighter and far more photo-stable.

Good luck!

AJH

 

Further reading