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A Better Method for Counting Brain Synapses

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Researchers at Washington University in St. Louis find an easier way to quantify synaptic loci

The brain’s vast synaptic network with billions of neurons making trillions of connections in a very dense matrix continues to challenge scientists who wish to decipher its intricacies in the context of brain health and disease. Established approaches using electron microscopy to visualize and study this elaborate landscape can be hugely time consuming and/or technically difficult to implement.

Andrew Sauerbeck and Terrance Kummer

Dr. Terrance Kummer (right) with Dr. Andrew Sauerbeck, first author of the article

Dr. Terrance Kummer and his team at the Hope Center for Neurological Disorders at Washington University in St. Louis (USA) have published an article describing a new method for quantifying brain synapses using ZEISS Airyscan technology. This method is simpler to implement and could enable more researchers to apply these types of analyses to critical questions in health and disease of the nervous system.

Dr. Kummer answered some questions for us about his research and this new method.

What are your research goals?

The overall goals of the Kummer lab are to understand how traumatic brain injury (TBI) damages the brain in the short term, and how these processes lead to long-term degeneration and cognitive decline.

TBI is the leading cause of death and disability for younger people, and also the best-established environmental risk factor for Alzheimer’s disease and related dementias. The connections between early brain trauma and later neurodegeneration, however, are not clear. We believe that fundamental processes likely underlie immediate disability after TBI and precipitate the development of dementia down the road. We hope to identify these processes, target them with specific therapies, and ultimately improve outcomes for the victims of TBI that I care for in the hospital.

Low-magnification image of brain mouse brain section labeled against the presynaptic marker synapsin (green) and the postsynaptic marker PSD-95 (magenta). Nuclei are blue. Image acquired with ZEISS Axio Scan.Z1 slide scanning system.

Low-magnification image of brain mouse brain section labeled against the presynaptic marker synapsin (green) and the postsynaptic marker PSD-95 (magenta). Nuclei are blue. Image acquired with ZEISS Axio Scan.Z1 slide scanner.

Why quantify brain synapses in order to study traumatic brain injury?

The highlighted publication reports on a method we devised to better characterize one possible source of short and long-term disability after TBI—synapse loss.

Heat map of synaptic density across the hippocampus of a control mouse (left) and an animal carrying a mutation that leads to deposition of amyloid plaques. Warmer colors denote greater density of synaptic loci.
Heat map of synaptic density across the hippocampus of a control mouse (left) and an animal carrying a mutation that leads to deposition of amyloid plaques. Warmer colors denote greater density of synaptic loci.

We were motivated to investigate synaptic injury after TBI because synapse loss is an early and prevalent finding in Alzheimer’s disease and many other dementing conditions. Synapse loss, moreover, is a better predictor of cognitive decline in Alzheimer’s disease than any other pathological feature, including plaques and tangles. We wondered whether TBI might induce synapse loss, thereby preventing neurons from communicating and increasing the risk of later dementia.

Tell us a little bit about your new method for analyzing brain synapses

Characterizing the structure and molecular features of synapses in the brain is challenging. Synapses are both below the resolution of traditional light microscopic techniques and packed at extreme densities in an extraordinarily complex milieu of cellular structures.

Pre- (green) and postsynaptic markers (red) are Airyscan imaged in the same field.

Pre- (green) and postsynaptic markers (red) are imaged with superresolution using ZEISS Airyscan.

Several approaches have been devised to overcome this problem, such as electron microscopy, array tomography, and optical super resolution, but all have significant drawbacks that limit their use.

We sought to develop a more accessible and straightforward method to identify, characterize, and quantify synapses that would help the approaches catch up to the field. That is, we wanted a technique that researchers could employ to answer questions involving many samples, many conditions, across age, and across space (cover large regions of the brain). All of these are necessary to understand brain injury and its link to long-term outcomes.

Video shows workflow for analyzing synaptic loci over large regions of brain using SEQUIN. Fully automated tile scanning is used to image very large regions, in this case an entire section of mouse hippocampus, at super-resolution (upper left). Individual images from this scan, when magnified, reveal individual pre- and postsynaptic puncta (lower left; green for pre- and magenta for postsynaptic puncta). SEQUIN analysis is used to measure pre-to-postsynaptic separations for individual puncta pairs, revealing a peak of puncta pairs consistent with synaptic loci (large peak in lower right graph). Quantifying this peak for individual parts of the tile scan allows us to build heat maps of synaptic density (upper right; warmer colors represent areas of greater synaptic density).

Why is it called “SEQUIN”?

We chose the name “SEQUIN” for the technique because it was both a good distillation of the key features of the analysis method–Synaptic Evaluation, QUantitation, and the images it produces reminded us of sequins glinting from a dark field. There was an active debate in the lab about what to call it! I prevailed, and everyone else pretended to be happy about it.

SEQUIN multiscale imaging seeks to understand brain structure and function by revealing the patterns of its synaptic connections, here represented by sequins, perhaps one day over the entire brain. Artwork credit: Story Kummer

SEQUIN multiscale imaging seeks to understand brain structure and function by revealing the patterns of its synaptic connections, here represented by sequins, perhaps one day over the entire brain. Artwork credit: Story Kummer

Where do you think this new method will bring an impact in research?

We believe that SEQUIN will open the door for non-specialist researchers to add synaptic structural and molecular analysis to their research questions. Said another way, we hope that labs that don’t focus full-time on synaptic analysis will now nonetheless have a low burden pathway to rigorously investigate synaptic endpoints in their studies. We believe that synaptic alterations, both adaptive and pathological, are an extremely common feature of the brain’s response to stimuli, whether those are developmental, experience-dependent, or the result of a brain injury or disease. We expect SEQUIN will enable a better understanding of these important processes.

Learn More

Read the full publication SEQUIN Multiscale Imaging of Mammalian Central Synapses Reveals Loss of Synaptic Connectivity Resulting from Diffuse Traumatic Brain Injury

Visit Dr. Kummer’s lab webpage or follow him on Twitter.

Get more information on ZEISS Airyscan, available on most ZEISS laser scanning confocal microscopes.

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The post A Better Method for Counting Brain Synapses appeared first on Microscopy.


Imaging Cilia and Centrioles Below the Diffraction Limit

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Novel use of expansion microscopy with ZEISS Airyscan technology

Primary cilia are organelles that protrude from the cell surface and function as “cellular antennae” to receive and transduce extracellular signals. They are approximately 3 µm in length and 0.3 µm in diameter, which is below the optical diffraction limit of standard light microscopes.

Junior Associate Professor Yohei Katoh (above, right) and Professor Kazuhisa Nakayama (above, left) at the Department of Physiological Chemistry, Kyoto University (Japan) have developed a novel technique to combine expansion microscopy (ExM) with ZEISS Airyscan superresolution microscopy technology to successfully observe cilia and centrioles with high brightness and resolution. Their findings are published here.

We spoke with Dr. Katoh about their work.

What are the research goals of your lab?

The Nakayama lab has been focusing on biogenesis of primary cilia and the mechanisms underlying regulation of ciliary protein trafficking.

The intraflagellar transport (IFT) machinery powered by kinesin and dynein motor proteins mediates anterograde and retrograde protein trafficking. The IFT machinery is a huge protein complex containing five multisubunit complexes composed of ~40 subunits in total. The Nakayama lab has revealed the architectures of these multisubunit complexes in the IFT machinery, the roles of individual subunits in the complexes, and the recognition mechanisms of cargo proteins by the complexes, and thereby elucidated the molecular mechanisms underlying the regulation of ciliary protein trafficking.

Primary cilia in hTERT-RPE1 cells

Primary cilia in hTERT-RPE1 cells imaged with conventional microscopy.

Defects in cilia biogenesis and ciliary protein trafficking therefore lead to various hereditary disorders with a broad spectrum of clinical manifestations, generally called the “ciliopathies.” The Nakayama lab also aims to understand the molecular and cellular basis of the ciliopathies.

What gave you the idea to combine Expansion microscopy (ExM) with Airyscan superresolution microscopy?

I had no experience with super-resolution microscopes, such as Airyscan or SIM (structured illumination microscopy), before starting this research. Since there is no super-resolution microscope in our institute as common equipment, I was looking for another method of super-resolution imaging of primary cilia.

When I first read the paper on expansion microscopy (ExM), I was impressed with the innovative idea of using superabsorbent gel to expand the cells. At that time, however, I wondered if this was really possible, and decided to try this innovative strategy myself. I bought some reagents, did some experiments, and was very surprised when I could really make cells expand.

Afterward, at a meeting of the Japanese Biochemical Society held in Osaka, I met Dr. Shuhei Chiba (Osaka City University), who is a skilled microscopist and utilizes super-resolution microscopy to study primary cilia and centrioles. At the meeting, Shuhei was very interested in my ExM images, and Shuhei and I came up with the idea to combine ExM and super-resolution microscopy techniques. I remember that I visited his lab with my ExM samples in the next week.

Dr. Shuhei Chiba. The monitor shows a primary cilium in a post-expansion sample.
Dr. Shuhei Chiba. The monitor shows a primary cilium in a post-expansion sample.
Airyscan analysis of the localization of MyosinVa (green; ciliary vesicle), polyglutamylated tubulin (pGlu-tubulin, red; axonemal microtubules), and CEP164 (blue; distal appendages of mother centriole) in non-expansion (left) and post-expansion (right) cells. The combinatorial use of Airyscan and expansion microscopy dramatically improves the optical resolution and makes multicolor super-resolution imaging more practical.
Airyscan analysis of the localization of MyosinVa (green; ciliary vesicle), polyglutamylated tubulin (pGlu-tubulin, red; axonemal microtubules), and CEP164 (blue; distal appendages of mother centriole) in non-expansion (left) and post-expansion (right) cells. The combinatorial use of Airyscan and expansion microscopy dramatically improves the optical resolution and makes multicolor super-resolution imaging more practical.

What was your reaction when you saw the data?

Of course, we were amazed at the image quality. Centrioles, which were only visible as dots under a conventional microscope, appeared to be beautiful nine-fold symmetrical structures when observed by the combinatorial use of ExM and Airyscan.

Airyscan image of the localization of intraflagellar transport protein 88 (IFT88), a subunit of anterograde IFT-B complex, and CEP164 in axially oriented mother centriole in post-expanded cells. The averaged maximum projection (AVG) images (bottom three panels) were generated by merging the set of nine-images obtained by rotating maximum intensity projection every 40° into one-stack.
Airyscan image of the localization of intraflagellar transport protein 88 (IFT88), a subunit of anterograde IFT-B complex, and CEP164 in axially oriented mother centriole in post-expanded cells. The averaged maximum projection (AVG) images (bottom three panels) were generated by merging the set of nine-images obtained by rotating maximum intensity projection every 40° into one-stack.

What do you plan to work on next?

The selective entry and exit of proteins into and out of cilia is controlled by the transition zone (TZ) located at the base of cilia. The TZ is composed of more than 20 different proteins, the mutations of which cause ciliopathies with a broad spectrum of clinical manifestations. However, little is known about how the TZ is constructed by these TZ proteins and how it controls the passage of ciliary proteins. Because of the limitations of standard light microscopy to observe the fine structure of the TZ, we are applying ExM and Airyscan to observe the TZ and to elucidate the molecular mechanisms underlying the selective entry and exit of ciliary proteins.

Learn More

Read the full article: Practical method for superresolution imaging of primary cilia and centrioles by expansion microscopy using an amplibody for fluorescence signal amplification Link

Learn about Expansion Microscopy

Get more information on Airyscan, a confocal detector which enables superresolution microscopy on new and many legacy ZEISS LSMs.

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The post Imaging Cilia and Centrioles Below the Diffraction Limit appeared first on Microscopy.

Microscopy at the MRC-University of Glasgow Centre for Virus Research

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Meet four scientists using different types of light microscopy for virology research

The MRC-University of Glasgow Centre for Virus Research (CVR) was established in 2010 and represents the UK’s largest grouping of human and veterinary virologists.

The CVR is embedded within the Institute of Infection, Immunity and Inflammation at the University of Glasgow, which provides excellent research opportunities to investigate virus-host interactions and immune response to virus infection. A defining feature of the CVR is its breadth of expertise ranging from molecular virology to in vivo pathogenesis, virus-cell interactions, viral immunology, viral ecology, viral oncology, clinical and veterinary virology, viral diagnostics, virus epidemiology, mathematical modelling, virus genomics & bioinformatics.

We profiled four scientists who use light microscopy to highlight some of the research being performed at the CVR.

Studying viruses with light sheet microscopy

Dr. Steph Rainey is a Research Fellow working within Professor Steven Sinkins’ Group.

She works on trying to understand how the bacterium Wolbachia blocks the replication of clinically important arboviruses, such as dengue. The primary vector for dengue, Aedes aegypti (mosquito), does not naturally harbor Wolbachia, which is found in ~66% of insects. However, the Sinkins’ group is able to transinfect the bacterium into the mosquito and have shown a 80% reduction in dengue cases in their Malaysian release site, compared to control sites.

Dr. Steph Rainey in front of her mosquito colonies
Dr. Steph Rainey in front of her mosquito colonies

To understand how Wolbachia blocks viruses, Dr. Rainey and the group uses a range of techniques to look at what is going on at a cellular level. They infect cells and mosquitoes with different arboviruses (arthropod/insect borne viruses) and then use microscopy to see how Wolbachia affects the replication complexes, where the virus multiplies and how both affect the structures inside the cell. To do this they use both confocal and light sheet microscopy. They use live samples, fixed samples, and cells.

One of the things they have found is that cholesterol trafficking is important for blocking viruses when Wolbachia is present. They also use fluorescence in situ hybridization to look at density and location of the Wolbachia within tissues and cells.

When asked why she likes using light sheet microscopy:

There is nothing like a good picture to tell a story. A lot of the time science is about numbers and graphs. I have a very visual brain so being able to see what is going on helps a lot.

Whole mosquitoes were cleared in hydrogen peroxide for four days. Images were obtained using a ZEISS light sheet microscope and rendered in Imaris. By simply rendering the autofluorescence we are able to see in amazing detail the anatomy of the head, mouth parts and eyes of the mosquito. Image shows four sides of one head.

Four sides of a mosquito head imaged using a ZEISS light sheet microscope and rendered in Imaris. We are able to see in amazing detail the anatomy of the head, mouth parts and eyes of the mosquito. Image credit: Stephanie Rainey, Maria Vittoria Mancini & Colin Loney

Superresolution imaging to better understand herpes simplex virus-1 (HSV-1)

Dr. Mila Collados Rodriguez, MRC-University of Glasgow Centre for Virus Research (CVR)

Dr. Mila Collados Rodriguez is a postdoctoral researcher working within Dr. Chris Boutell’s Group. Currently, she aims to determine how molecular components of promyelocytic leukaemia-nuclear bodies (PML-NBs) drive the repression of the herpes simplex virus-1 (HSV-1) genome.

Dr. Collados Rodriguez uses the Airyscan detector of a ZEISS confocal microscope for superresolution imaging.

Cells are seeded onto cover slips which are either mock treated or infected with labelled HSV-1. At different time-points, cells are permeabilized, fixed and then conjugated with a fluorophore (click chemistry) which allows visualization of the labelled viral DNA. Immunofluorescence on PML-NB components is also performed to investigate the spatiotemporal contribution of PML-NB molecules in restricting HSV-1 infection at a single viral genome resolution.

Left, ZEISS LSM 880 Airyscan 2D projection of a human fibroblast cell nucleus (stained with DAPI, blue) harboring a click chemistry red labelled HSV-1 genome further detailed in 3D. Right, same image evidencing colocalization and entrapment of the HSV-1 genome by PML immunostaining (cyan). Mila Collados Rodríguez, CVR-University of Glasgow, UK.

Left, ZEISS LSM 880 Airyscan 2D projection of a human fibroblast cell nucleus (stained with DAPI, blue) harboring a click chemistry red labelled HSV-1 genome further detailed in 3D. Right, same image evidencing colocalization and entrapment of the HSV-1 genome by PML (cyan) immunostaining. Mila Collados Rodríguez, CVR-University of Glasgow, UK

Cell nucleus containing 2 HSV-1 genomes (labelled in red) entrapped by PML (in cyan) imaged in 3D with ZEISS LSM 880 with Airyscan and processed with IMARIS software. Mila Collados Rodríguez, CVR-University of Glasgow, UK.

Using live cell imaging to visualize coinfection of viruses

Joanne Haney is a PhD student working within Dr. Pablo Murcia’s Group. She studies virus interactions during co-infection between common respiratory viruses. She is interested in the dynamics of infection when two viruses are replicating in the same population of cells. She compares the phenotypes of infection between each virus replicating alone, and in the presence of one another to identify differences that may highlight virus-virus interactions.

Live cell imaging is a huge part of her research.

Joanne Haney, MRC-University of Glasgow Centre for Virus Research (CVR)

She is able to observe how viral infections spread throughout a population of cells and what effect this has on cell survival and cell movement. It also allowed her to track individual cells to determine how the infection status of that cell (infected with one or both viruses) is related to the viral induced outcome.

Human lung cells infected with two seasonal respiratory viruses that have been tagged with fluorescent proteins. Cell nuclei were stained (blue) to allow them to be easily tracked and cells were imaged over a time course to capture the dynamics of infection and viral spread.

Single cell analysis using laser microdissection to better understand hepatitis C virus

Carol Leitch, MRC-University of Glasgow Centre for Virus Research

Dr. Carol Leitch is a Research Fellow working within Professor John McLauchlan’s Group. She researches how hepatitis C virus (HCV) affects liver cells (hepatocytes) by comparing the transcriptomes of individual infected and uninfected cells. The end goal of her research is to be able to describe the effect of HCV on both infected and uninfected hepatocytes spatially at the single cell level.

Dr. Leitch uses a laser microdissection microscope, which is extremely accurate, to excise single cells from liver biopsy specimens. To prepare the samples, she uses a cryomicrotome to section fresh-frozen liver needle biopsies. The sections are then fixed in formaldehyde and RNA-FISH is performed with multiple probes against the HCV RNA genome to identify infected cells.

Read select publications from the CVR:

  • The Wolbachia strain wAu provides highly efficient virus transmission blocking in Aedes aegypti. PLOS Pathogens
  • Distinct temporal roles for the promyelocytic leukaemia (PML) protein in the sequential regulation of intracellular host immunity to HSV-1 infection. PLOS Pathogens
  • Perturbed cholesterol and vesicular trafficking associated with dengue blocking in Wolbachia-infected Aedes aegypti cells. Nature Communications

Light microscopy technology referenced in this article:

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The post Microscopy at the MRC-University of Glasgow Centre for Virus Research appeared first on Microscopy.

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