この作品は、NIHのグラントEY 14990によってサポートされていました。

<ol>
	<li>Axelrod, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Total+internal+reflection+fluorescence+microscopy+in+cell+biology.">Total internal reflection fluorescence microscopy in cell biology.</a> Traffic. 2, 764-774 (2001).</li><li>Zenisek, D., Steyer, J. A., Almers, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport,+capture+and+exocytosis+of+single+synaptic+vesicles+at+active+zones.">Transport, capture and exocytosis of single synaptic vesicles at active zones.</a> Nature. 406, 849-854 (2000).</li><li>Zenisek, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vesicle+association+and+exocytosis+at+ribbon+and+extraribbon+sites+in+retinal+bipolar+cell+presynaptic+terminals.">Vesicle association and exocytosis at ribbon and extraribbon sites in retinal bipolar cell presynaptic terminals.</a> Proc. Natl. Acad. Sci. U. S. A. 105, 4922-4927 (2008).</li><li>Zenisek, D., Horst, N. K., Merrifield, C., Sterling, P., Matthews, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+synaptic+ribbons+in+the+living+cell.">Visualizing synaptic ribbons in the living cell.</a> J. Neurosci. 24, 9752-9759 (2004).</li><li>Axelrod, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+imaging+of+surface+fluorescence+with+very+high+aperture+microscope+objectives.">Selective imaging of surface fluorescence with very high aperture microscope objectives.</a> J. Biomed. Opt. 6, 6-13 (2001).</li><li>Rouze, N. C., Schwartz, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continous+and+transient+vesicle+cycling+at+a+ribbon+synapse.">Continous and transient vesicle cycling at a ribbon synapse.</a> J. Neurosci. 18, 8614-8624 (1998).</li></ol>

客観型全反射顕微鏡の利点は、1）それはそれによって焦点のずれた光を最小限に抑え、目的の焦点面内に狭い領域に励起光を制限することにより、優れた光学セクショニングを提供することであり、2）光が距離と共に指数関数的に低下するので、 、垂直方向の移動は、蛍光強度の変化としてモニターすることができる、高開口数の対物1,5〜3）効率的な集光。 技法の主な欠点は、それが電子?...

imaging exocytosis in retinal bipolar cells with tirf microscopy

全内部反射蛍光（全反射）顕微鏡は、ガラスなどの高屈折率物質に最も近い蛍光分子の選択的なイメージングによって、細胞膜で発生するイベントの調査を可能にする技術です。<sup&gt; 1</sup&gt;。この記事では、金魚の網膜から単離された網膜双極細胞におけるシナプス小胞の画像のエキソサイトーシスにこの手法を適用する。これらのニューロンは、彼らの大きな軸索末端に起因するこの種の研究に非常に適している。双極細胞のクランプを同時にパッチすることで、それはシナプス前電圧とシナプス放出との関係を調査することが可能です。<sup&gt; 2,3</sup&gt;。双極細胞の端末内のシナプス小胞は、共同パフ色素と高Kを含むリンゲル液によって蛍光染料（FM 1から43 ®）を使用してロードされます<sup&gt; +</supシナプス端子へ&gt;濃度。これは、細胞を脱分極し、グルタミン酸作動性小胞へのエンドサイトーシスとそれに伴う色素の取り込みを刺激する。約30分間離れて余分な色素を洗浄後、細胞はパッチがクランプと488 nmのレーザーで同時に撮像されるための準備が整いました。パッチピペット溶液は、シナプスリボンのタンパク質のリブアイに選択的に結合するローダミンベースのペプチドが含まれています<sup&gt; 4</sup&gt;、端末が561 nmのレーザーで撮像する場合、それによって具体的にリボンにラベルを付ける。これは、アクティブなゾーンの正確な局在と余分なシナプスのイベントからシナプスを分離することができます。

Watch this Scientific Journal Video about Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy at JoVE.com

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

このビデオでは、我々は内部全反射蛍光（全反射）顕微鏡を使用して金魚網膜双極細胞における単一シナプス小胞のエキソサイトーシスと人身売買にラベルを付けたり、視覚化する方法を示しています。

全反射顕微鏡による網膜双極細胞におけるイメージングのエキソサイトーシス

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

imaging-exocytosis-in-retinal-bipolar-cells-with-tirf-microscopy

Research

JoVE Journal

Biology

11.6K ビュー.  Yale University School of Medicine. このビデオでは、我々は内部全反射蛍光（全反射）顕微鏡を使用して金魚網膜双極細胞における単一シナプス小胞のエキソサイトーシスと人身売買にラベルを付けたり、視覚化する方法を示しています。

ビデオ: Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

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.

広視野顕微鏡を用いて長期的なイメージング哺乳類細胞

生物学

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

光イメージングによる非侵襲的検出のための蛍光色素で標識幹細胞

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

siRNAトランスフェクション後の有糸分裂のタイムラプスイメージング

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

生細胞におけるストア作動性カルシウムエントリの蛍光ベースの測定：培養癌細胞から骨格筋線維に

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

成体マウス頭蓋冠骨髄中の造血幹細胞および前駆細胞のin vivo 4次元トラッキングで

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

ワンナノメートルの精度で蛍光イメージング（FIONA）

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

タンデム高圧凍結および透過型電子顕微鏡のための植物組織の急速凍結置換

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

動的細胞突起におけるソフトウェア支援トラッキングのためのグラフィカルユーザインタフェース

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated exocytosis is fundamental for intercellular communication and is a key mechanism for the secretion of neurotransmitters, hormones, inflammatory mediators, and other compounds, by a variety of cells. At least three distinct mechanisms are known for regulated exocytosis: full exocytosis, where a single SG fully fuses with the plasma membrane, kiss-and-run exocytosis, where a single SG transiently fuses with the plasma membrane, and compound exocytosis, where several SGs fuse with each other, prior to or after SG fusion with the plasma membrane. The type of regulated exocytosis undertaken by a cell is often dictated by the type of secretory trigger. However, in many cells, a single secretory trigger can activate multiple modes of regulated exocytosis simultaneously. Despite their abundance and importance across cell types and species, the mechanisms that determine the different modes of secretion are largely unresolved. One of the main challenges in investigating the different modes of regulated exocytosis, is the difficulty in distinguishing between them as well as exploring them separately. Here we describe the use of fluorescein isothiocyanate (FITC)-dextran as an exocytosis reporter, and live cell imaging, to differentiate between the different pathways of regulated exocytosis, focusing on compound exocytosis, based on the robustness and duration of the exocytic events.

Here we detail a method for live cell imaging of regulated exocytosis. This method utilizes FITC-dextran, which accumulates in lysosome-related organelles, as a reporter. This simple method also allows distinguishing between different modes of regulated exocytosis in cells that are difficult to manipulate genetically.

規制エキソサイトーシスの記者として FITC デキストランをイメージング

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated ...