This work was supported by a grant from the AHA to R.E.D. (15SDG25560035). Authors would like to acknowledge Dr. Fernando Santana for use of his Leica SR GSD 3D microscope, and Dr. Johannes Hell for kindly providing the FP1 antibody.

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The authors have no competing interests to disclose.

The recent explosion of technologies that allow imaging beyond the diffraction limit have offered new windows into the complexities of mammalian cell signaling in space and time. These technologies include STORM, STED, PALM, GSD, SIM, and their variants (e.g., dSTORM, FPALM). The ingenuity of the scientists behind these techniques has allowed us to circumvent the limitations imposed by laws of physics governing the diffraction of light. In spite of this huge accomplishment, each of these techniques makes some so...

Cellular signaling reactions translate changing internal and external environments to initiate a cellular response. They regulate all aspects of human physiology, serving as the foundation for hormone and neurotransmitter release, the heartbeat, vision, fertilization, and cognitive function. Disruption of these signaling cascades can have severe consequences in the form of pathophysiological conditions including cancer, Parkinson&#39;s, and Alzheimer&#39;s disease. For decades, biological and medical investigators have successfully used fluorescent proteins, probes, and biosensors coupled with fluorescence microscopy as the primary tools to understand the precise spatial and temporal organization of these cellular signals.
The strengths of optical techniques such as epifluorescence, confocal, or total internal reflection fluorescence (TIRF) microscopy are their sensitivity, speed, and compatibility with live cell imaging, while the major limitation is their diffraction-limited resolution, meaning structures or protein complexes smaller than 200-250 nm cannot be resolved. With the theoretical and practical development of deterministic super-resolution (e.g., stimulated emission depletion microscopy (STED1), structured illumination microscopy (SIM2) or stochastic super-resolution (e.g., photoactivated localization microscopy (PALM3), or ground state depletion (GSD4,5)), lateral and axial resolution in fluorescence microscopy has been extended beyond the diffraction barrier, to the order of tens of nanometers. Thus, investigators now have the unparalleled ability to visualize and understand how protein dynamics and organization translates to function at the near-molecular level.
Ground state depletion microscopy followed by individual molecule return (GSDIM), or simply GSD as it is known, circumvents the diffraction limit by reducing the number of simultaneously emitting fluorophores4,5. High energy laser light is used to excite the fluorophore-labelled sample, bombarding electrons with photons and increasing the probability they will undergo a &#39;spin-flip&#39; and enter the triplet or &#39;dark-state&#39; from the excited state4. This effectively depletes the ground state, hence the name &#39;ground state depletion&#39;. In the triplet state, fluorophores do not emit photons and the sample appears dimmer. However, these fluorophores stochastically return to the ground state and can go through several photon emitting excited-to-ground state transitions before returning to the triplet state. With less fluorophores emitting at any given time, photon bursts emitted from individual fluorophores become spatially and temporally distinct from neighboring fluorophores. The burst of photons can be fit with a gaussian function, the calculated centroid of which corresponds to the position of the fluorophore with a localization precision that is dependent on the numerical aperture (NA) of the lens, the wavelength of light used for excitation and crucially, the number of photons emitted per fluorophore. One limitation of GSD is that, since only a subset of fluorophores actively emits at any time, thousands of images must be collected over several minutes to build up a complete localization map. The long acquisition time combined with the high laser power requirement, means that GSD is better suited to fixed rather than live samples.
This article, describes the preparation of fixed samples for super-resolution microscopy imaging of membrane and endoplasmic reticulum (ER)-resident proteins&#160;(for a list of necessary consumables and reagents see the Table of Materials). Examples of how this protocol can be easily adapted to quantify the size and degree of clustering of L-type voltage-gated Ca2+ channels (Cav1.2) in the sarcolemma of cardiac myocytes, or used to visualize the cellular distribution of the ER, are demonstrated. Understanding the distribution and organization of these cellular components is critically important in understanding the initiation, translation, and ultimately the function of many Ca2+-dependent signaling cascades. For example, Cav1.2 channels are fundamentally important for excitation-contraction coupling, while receptor-mediated Ca2+ release from the ER is perhaps the most ubiquitous signaling cascade in mammalian cells.

<table><tbody><tr><td>KOH</td><td>Thermo Scientific</td><td>P250-500</td><td>To clean coverglass</td></tr><tr><td>#1.5 coverglass 18 x 18 mm</td><td>Marienfeld Superior</td><td>0107032)</td><td>To grow/process/image cells</td></tr><tr><td>10x PBS</td><td>Thermo Scientific</td><td>BP3994</td><td>Dilute to 1x with de-ionized water</td></tr><tr><td>Poly-L-Lysine</td><td>Sigma</td><td>P4832</td><td>Aids with cell adhesion to cover glass</td></tr><tr><td>laminin</td><td>Sigma</td><td>114956-81-9</td><td>Aids with cell adhesion to cover glass</td></tr><tr><td>Medium 199</td><td>Thermo Scientific</td><td>11150-059</td><td>Ventricular myocyte culture media</td></tr><tr><td>DMEM 11995</td><td>Gibco</td><td>11995</td><td>Cell culture media</td></tr><tr><td>Fetal bovine Serum (FBS)</td><td>Thermo Scientific</td><td>10437028</td><td>Media supplement</td></tr><tr><td>Penicillin/streptomycin</td><td>Sigma</td><td>P4333</td><td>Media supplement</td></tr><tr><td>0.05% trypsin-EDTA</td><td>Corning</td><td>25-052-CL</td><td>Cell culture solution</td></tr><tr><td>Lipofectamine 2000</td><td>Invitrogen</td><td>11668-019</td><td>Transient transfection reagent</td></tr><tr><td>Ca2+-free PBS</td><td>Gibco</td><td>1419-144</td><td>Cell culture</td></tr><tr><td>100 % Methanol</td><td>Thermo Fisher Scientific</td><td>A414-4</td><td>Cell Fixation</td></tr><tr><td>Paraformaldehyde</td><td>Electron Microscopy Sciences</td><td>15710</td><td>Cell Fixation</td></tr><tr><td>Glutaraldehyde</td><td>Sigma Aldrich</td><td>SLBR6504V</td><td>Cell Fixation</td></tr><tr><td>SEAblock</td><td>Thermo Scientific</td><td>37527</td><td>BSA or other blocking solution alternatives exist</td></tr><tr><td>Triton-X 100</td><td>Sigma</td><td>T8787</td><td>Detergent to permeabilize cells</td></tr><tr><td>Rabbit anti-CaV1.2 (FP1)</td><td>Gift</td><td>N/A</td><td>Commercial anti-CaV1.2 antibodies exist such as Alomone Labs Rb anti-CaV1.2 (ACC-003)</td></tr><tr><td>Mouse monoclonal anti-RFP</td><td>Rockland Inc.</td><td>200-301-379</td><td>Primary antibody</td></tr><tr><td>Alexa Fluor 647 donket anti-rabbit IgG (H+L)</td><td>Invitrogen (Thermo Scientific)</td><td>A31573</td><td>Secondary antibody</td></tr><tr><td>Alexa Fluor 568 goat anti-mouse IgG (H+L)</td><td>Invitrogen (Thermo Scientific)</td><td>A11031</td><td>Secondary antibody</td></tr><tr><td>Sodium azide</td><td>Sigma</td><td>S2002</td><td>Prevents microbial growth for long term storage of samples</td></tr><tr><td>Catalase</td><td>Sigma</td><td>C40</td><td>Photoswitching buffer ingredient</td></tr><tr><td>Glucose oxidase</td><td>Sigma</td><td>G2133</td><td>Photoswitching buffer ingredient</td></tr><tr><td>Tris</td><td>Sigma</td><td>T6066</td><td>Photoswitching buffer ingredient</td></tr><tr><td>beta-Mercaptoethylamin hydrochloride</td><td>Fisher</td><td>BP2664100</td><td>Photoswitching buffer ingredient</td></tr><tr><td>&amp;beta;-mercaptoethanol</td><td>Sigma</td><td>63689</td><td>Photoswitching buffer ingredient</td></tr><tr><td>NaCl</td><td>Fisher</td><td>S271-3</td><td>Photoswitching buffer ingredient</td></tr><tr><td>Dextrose</td><td>Fisher</td><td>D14-212</td><td>Photoswitching buffer ingredient</td></tr><tr><td>Glass Depression slides</td><td>Neolab</td><td>1 &amp;ndash; 6293</td><td>To mount samples</td></tr><tr><td>Twinsil</td><td>Picodent</td><td>13001000</td><td>To seal coverglass</td></tr><tr><td>sec61&amp;beta;-mCherry plasmid</td><td>Addgene</td><td>49155</td><td></td></tr><tr><td>Leica SR GSD 3D microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>ImageJ</td><td></td><td></td><td></td></tr><tr><td>Washing block solution</td><td></td><td></td><td>20 % SEAblock in PBS</td></tr><tr><td>Primary antibody incubation solution</td><td></td><td></td><td>0.5 % Triton-X100, 20 % SEAblock, in PBS</td></tr><tr><td>Secondary antibody incubation solution</td><td></td><td></td><td>1:1000 in PBS</td></tr></tbody></table>

ground state depletion super-resolution imaging in mammalian cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

As documented in the introduction, there are many different super-resolution microscopy imaging modalities. This protocol, focuses on GSD super-resolution imaging. Representative images and localization maps are shown in Figure 2 and Figure 3.
Figure 2 shows a COS-7 cell transfected with the ER protein, mCherry-Sec61&#946;, and processed us...

Watch this Scientific Journal Video about Ground State Depletion Super-resolution Imaging in Mammalian Cells at JoVE.com

Ground State Depletion Super-resolution Imaging in Mammalian Cells

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

ground-state-depletion-super-resolution-imaging-in-mammalian-cells

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7.2K Views.  University of California School of Medicine, Davis. Dit artikel schetst een protocol voor de detectie van een of meer plasma en/of intracellulaire membraaneiwitten grondtoestand uitputting (GSD) Super resolutie microscopie met zoogdiercellen. Hier bespreken we de voordelen en de overwegingen van het gebruik van dergelijke benaderingen voor de visualisatie en de kwantificering van cellulaire eiwitten.

Video: Ground State Depletion Super-resolution Imaging in Mammalian Cells

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Actually what I'm gonna do, mount it in here first and then swap out You. So now I'm gonna aspirate try and be as gentle as possible.Okay. A few seconds just holding.Okay.Alright.So the important features of our lifestyle imaging grid are that, so it's an inverted scope, which means the objective, the samples here in the objective is underneath. So the objective is pointing up and the whole scope, or practically the whole, practically the whole scope, including the objectives and the sample are at 37 degrees. So everything in this plastic box is at 37 degrees.And right on top of the stage we have an air and CO2 mix. That is 5%CO2. So the idea is that the sample is actually in a cell, it's like a cell incubator.And so we have, this is our mixer for the air and the CO2 and, and it gets humidified and heated and then piped onto the, the sample on the stage.Okay. And today we're just gonna be doing fluorescence one channel GFP fluorescence. This is the hose that the CO2 and 5%CO2 goes in and it just fits into the stage holder.That's important. And then we can close these, take these down, and then we go to our first position. And what I'm gonna do is go to the first field, which is my control sample, and I'll pick positions based on what I see on the screen.So I just make sure I actually look into the microscope in each new, well, because the focal plane for each of the wells is different, the, the dish is not perfectly flat on the bottom. So you will need to make fine adjustments when you move from well to the well. So right now I'm on the first, well, what I'll do is I'll focus, so I just hit, I just told, I just turned on the light, the transmitted light for the microscope.And I have make sure all the light is going to the eye pieces. And so I just look in, make sure I get a nice clean field. And then what I'll do is turn that light off, turn the fluorescence light on, make sure the cells that I'm looking at look healthy and they're not apoptotic.And, and I turn that off. And what I do next is send, so now I'm gonna start picking positions, which means I have to te send the light path to the camera. This is the camera right here.So I tell the microscope to send all the camera, they don't see anything in the eye pieces. What I'll do is in the software controlling the microscope, I'll choose the right filter set, which for our purposes is psy. And then set an exposure time.And I know these cells very well and I know that they take a 0.2 second exposure time. So I tell, that's what I tell the software to do. And what I'll do is hit focus.This is, and it's basically just alive. The, A light will turn on and you can, oh, I don't know if you can see cells, but the reason this is a good field is because there's, all the cells are, even in terms of their fluorescence, There are no apoptotic cells. Apoptotic cells tend to be very bright and there's no dirt or schmutz in the field.And you can see that they're actually, this is a metaphase cell and that's a metaphase cell, that's a cell that's probably just exited mitosis. And I can just tell that because it looks like those are, those two sets of chromatin are very nicely oriented to each other. Like they just exited an face.So that's a good field. So, and now that I know that the exposure time and the filter set is correct, and this is gonna be software specific, I'll go and I'll take positions and now the software has memorized the X, Y, Z position of that field. And then when I do pick another field, it's, and I'll, so when I'm picking, when I'm picking new fields, but in the same, well, I'll use the, I'll move the i'll, I'll turn the shutter, I'll open the shutter and I'll look to see what I see on the screen and I'll pick fields based on that versus looking in the field.So I don't see any of my tonics in there. But again, even nicely centered, these cells will go through my PS I continue, here's a good example. Here's a good example of what I would avoid that, see how bright that is?That's very bright. That's probably remnants of an apoptotic cell. And the reason I would avoid that is because since it's so bright, when the software auto focuses, it will autofocus just on that and it won't be in the same plane as all these other cells.And so I won't get any information from that field. That looks nice. There's a, this is a pro metaphase right here, That's A metaphase.So, and I'm gonna take three positions per well. So that was three positions and now I'm going to go to my next Well, so you can, so what I will probably do is take images every 10 minutes. So it's a compromise.Lifestyle imaging is always a compromise, right? You have to image, you have to keep the cells healthy, especially if you're interested in mitosis. So you have to minimize their exposure to the fluorescent line.And, but you wanna get information. So you have to take images, you have to, so you have to compromise, you have to make a choice about what you want to capture, what kind of information you wanna capture. For our purposes, mitosis tends to take about an hour.So if we're taking 10 minute intervals, that's pretty good. We see prophase, we see metaphase, we see anaphase, and we can, we can see when there are, there are treatments or drugs or siRNAs that affect those specific cell cycle stages. So for our purposes, that's enough, but we always want more.We always want more positions or finer time and interval or something like that. So it's a compromise. So, and then I'll hit start.And so our software, it does an autofocusing pass on the first pass. So once I hit start, it should start auto autofocusing and you'll see the shutter open, open and close between auto focusing. So I'll hit start.So that's the shutter opening and closing. The Z motor is engaging and it's actually trying to find the plane with the most contrast, which will be the most in focus. You can actually see on the screen going in and out of finding the best plane of focus that might be worrisome.So what it does is it, it drives the Z motor to find the best, the most in focus plane focus, well the most in image. And then it acquires picture there and then it learns that Z position. And then we'll use that Z position for the next 12 passes.And then we'll do the auto focusing pass again. What Kind of experiments do they do? Okay, so our lab is mostly interested in understanding what regulates mitic progression, or at least I'm most interested in understanding that.So with live cell imaging and especially our multi position, long-term imaging scope, you can take, you can film cells for two or three days entering anding mitosis, and you can image multiple rounds of mitosis so you can understand. And then you can put drugs or RNAs or any kind of perturbations on the cells that you would like to study. So you can understand how, how those drugs or, or genes are, are important in regulating mitotic progression.Because it's not fixed cell work and live cell work, you can understand what, when you need to have a perturbation to affect mitosis or what stage of mitosis is affected. So it's really powerful in that respect. What, what are the main technical elements of this experiments?What are the possible problems? Well, the biggest problem besides getting the rig set up, getting the rig set up is not trivial. A lot of people have the rig, but just making sure it works well for your application.The financial element is not, you know, is not small. So there's the setting up of the apparatus and then, and then once you gather the data, there's a real data analysis issue. It takes these movie, if, so, if you're gathering 40 movies each with 30 cells, so you have 40 in 40 fields that you just filmed and each of those fields has 30 cells, then you have to sit down with each one of those movies and look at each one of those cells in that movie.So it takes a really long time to do data analysis. And that's, so it takes a week to analyze data that it took you two days to collect. So there's a, that's a very big obstacle in terms of gathering lots of data.So it's not, while it's high throughput data collection, it's not high throughput analysis. So what would you make to make this experiment really high throughput? So our lab is in the process of developing a program that does the data analysis automatically.And it's a pretty decent program. It's not perfect. The human eye is always better, but it's getting there.So What are the interesting application of this experiment? Maybe other fields you can mention? Besides cell division?You mean for the, oh, for the live cell imaging? Yes.Well, you can do cell motility assays. You can, depending on the, the microscopy that's involved, you can, you know, look at how cells, you could look at tumor development if you really wanted to.But there, you, you're, if as long as you have the right imaging set up, you can, you're pretty much not limited in terms of anything. Our scope happens to be just a wide field and we use 20 by 20 x objective. So it's a pretty low resolution set up, but that's sufficient for what we need to see.So there are, but there definitely aren't, you wouldn't be limited. Anything that has any dynamic kind of experiment, which is biology, it would be, would really benefit from lifestyle imaging. So calcium imaging, although you couldn't do it on our apparatus, that's a live cell.You definitely need live cell imaging for that. Like I said, patient assays, motility assays, those kinds of things would really benefit from live cell imaging. Would you like to tell a few words about your previous experience?How long do you work on these techniques? What do you you work on before? Wow.So in my PhD I didn't work.I did a lot of confocal microscopy, but no lifestyle imaging. And so during my postdoc I was, I helped Dr.King get this apparatus that, that we just used to set up these fragment, get that up and running. And, and so during my postdoc, I had a lot of experience with both widefield with TimeLapse imaging on our scope, which is the widefield and also TimeLapse imaging with confocal microscopy.And I would say that it's really powerful. You get a lot of information, but it's not easy to get working perfectly. So it really requires a lot of attention and a lot of attention to details at every little step of the process.And, and then you still have problems. So it's a tough application, but you can really get, if, if it's what you need to have to, to study your problem, your scientific problem, then it, it's invaluable in that respect, but it's tough. So find someone who knows what they're doing and use them.That's my advice. So good.

Conventional BODIPY Conjugates for Live-Cell Super-Resolution Microscopy and Single-Molecule Tracking

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve intracellular structures and the dynamics of biomolecules with ~20 nm precision. A prerequisite for SMLM are fluorophores that transition from a dark to a fluorescent state in order to avoid spatio-temporal overlap of their point spread functions in each of the thousands of data acquisition frames. BODIPYs are well-established dyes with numerous conjugates used in conventional microscopy. The transient formation of red-shifted BODIPY ground-state dimers (DII) results in bright single molecule emission enabling single molecule localization microscopy (SMLM). Here we present a simple but versatile protocol for SMLM with conventional BODIPY conjugates in living yeast and mammalian cells. This procedure can be used to acquire super-resolution images and to track single BODIPY-DII states to extract spatio-temporal information of BODIPY conjugates. We apply this procedure to resolve lipid droplets (LDs), fatty acids, and lysosomes in living yeast and mammalian cells at the nanoscopic length scale. Furthermore, we demonstrate the multi-color imaging capability with BODIPY dyes when used in conjunction with other fluorescent&#160;probes. Our representative results show the differential spatial distribution and mobility of BODIPY-fatty acids and neutral lipids in yeast under fed and fasted conditions. This optimized protocol for SMLM can be used with hundreds of commercially available BODIPY conjugates and is a useful resource to study biological processes at the nanoscale far beyond the applications of this work.&#160;

Conventional BODIPY conjugates can be used for live-cell single-molecule localization microscopy (SMLM) through exploitation of their transiently forming, red-shifted ground state dimers. We present an optimized SMLM protocol to track and resolve subcellular neutral lipids and fatty acids in living mammalian and yeast cells at the nanoscopic length scale.

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve ...

Biochemistry

Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has been in the study of immunological synapse formation, due to the ability of the glass-supported planar lipid bilayers to mimic the surface of a target cell while forming a horizontal interface. The recent advances in super-resolution imaging have further allowed scientists to better view the fine details of synapse structure. In this study, one of these advanced techniques, stimulated emission depletion (STED), is utilized to study the structure of natural killer (NK) cell synapses on the supported lipid bilayer. Provided herein is an easy-to-follow protocol detailing: how to prepare raw synthetic phospholipids for use in synthesizing glass-supported bilayers; how to determine how densely protein of a given concentration occupies the bilayer's attachment sites; how to construct a supported lipid bilayer containing antibodies against NK cell activating receptor CD16; and finally, how to image human NK cells on this bilayer using STED super-resolution microscopy, with a focus on distribution of perforin positive lytic granules and filamentous actin at NK synapses. Thus, combining the glass-supported planar lipid bilayer system with STED technique, we demonstrate the feasibility and application of this combined technique, as well as intracellular structures at NK immunological synapse with super-resolution.

We describe here a combination of the glass-supported lipid bilayer technique of forming immunological synapses with the super-resolution imaging technique of stimulated emission depletion (STED) microscopy. The goal of this protocol is to provide users with the instructions necessary to successfully carry out these two techniques.

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has ...

Immunology and Infection

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

Neuroscience

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Developmental Biology

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Super-Resolution Imaging to Study Co-Localization of Proteins and Synaptic Markers in Primary Neurons

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. Imaging studies are widely used to investigate their function and plasticity in physiological and pathological conditions. Because of their size and structure, localization studies of proteins require high-resolution imaging techniques. In this protocol, we describe a procedure to study in primary neurons the co-localization of target proteins with synaptic markers at a super-resolution level using structured illumination microscopy (SIM). SIM is a patterned-light illumination technique that doubles the spatial resolution of wide-field microscopy, reaching a detail of around 100 nm. The protocol indicates the required controls and settings for robust co-localization studies and an overview of the statistical methods to analyze the imaging data properly.

This protocol shows how to employ super-resolution microscopy to study protein co-localization in primary neuronal cultures.

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

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