Calcium Imaging - Science method

Sundas Arshad Try adjusting the following factors one at a time to see which has the most positive effect on your imaging results.

1. Ensure that Fluo-4 is loaded effectively: Optimise loading time, dye concentration and wash steps to avoid excessive background fluorescence that could mask the calcium spikes.

2. The time resolution of 1-2 s may be too slow to capture fast calcium spikes, especially if they are transient. Reducing the interval to 0.5s or even faster may help to capture transient events. However, be aware of increased photobleaching and phototoxicity and find a balance that works for your cells.

3.ATP at 2 µM may not be sufficient to elicit a clear calcium response in all cells, especially if receptor sensitivity varies between cells; consider increasing the ATP concentration stepwise to determine the optimum level for consistent responses. Also ensure that receptors for ATP (such as P2Y or P2X receptors) are expressed equally on all cells and that their activity is influenced by the state of differentiation.

4. LUHMES cells, even differentiated ones, can have different responses depending on their maturity and individual receptor expression. Ensure consistent differentiation of all cells (e.g. by optimising differentiation time or using specific markers to assess maturity) to reduce variability in response.

5. You also mentioned that problems with saline and DPBS cause cell stress, but the differentiation medium seems to maintain cell viability better. The composition of the imaging medium, such as ion concentrations (e.g. calcium and magnesium) and pH buffering capacity, can significantly affect cell response. The differentiation medium is likely to provide growth factors or supplements that help keep the cells healthy during imaging. You may wish to try a modified imaging buffer that mimics the composition of your differentiation medium, but is more suitable for fluorescence imaging. FluoroBrite DMEM or BrainPhys Imaging Buffer could be good options as they provide the necessary nutrients with minimal background fluorescence. Alternatively, you could use HBSS with supplements such as glucose and calcium to better mimic the conditions of the differentiation medium.

6. Finally, if you notice a general increase in calcium levels without oscillations, the cells may be under some kind of stress (possibly metabolic or mechanical). Ensuring gentle handling, using a low-power objective (20x is reasonable, but reducing the laser intensity might help), and minimising time out of the incubator might improve consistency.

I hope these suggestions help to optimise your calcium imaging experiments.

Hey Bipin Raj Shekhar ,

So, it is cellSens that you're using, am I correct? If so, check this tutorial, it may be helpful:

Also, keep in mind that if you're using it for your analysis, the software can be purchased in different packages (from the simplest to a full version). Know which package you have so you'll have access (or not) to all its functionalities.

Good luck,

Andre

I am not 100% clear on what you are asking, so forgive me if I'm answering things that are beside the point.

First of all, do you mean fluo 4, the calcium-sensitive indicator? This dye is best for measuring relative changes in calcium, and not as good for measuring absolute calcium levels. If you want to measure resting calcium inside cells, you should probably use a ratiometric dye such as fura-2. Grynkiewicz et al 1985 provides a good roadmap for how to quantify Ca. It is possible to do with a non-ratiometric dye like fluo-4, but it is more difficult. (See Brenowitz & Regehr 2003 for methodology.)

I'm not sure in your system what would constitute increased or decreased excitation. Differences in resting calcium are not commonly used as a readout of excitation. Rather, people often quantify action potential number or frequency. You can assess that sort of activity using calcium imaging, in which case a ratiometric dye isn't necessary, and fluo-4 is a good choice.

For "confocal parameters", I would say that things like gain and dwell time and resolution and stuff are whatever it takes to get a good, unsaturated image. By "unsaturated", I mean you want to be able to detect increases and decreases in brightness, so you don't want pixels to be at the maximum or minimum brightness values.

For assessing activity associated with action potentials, you would want to monitor fluorescence over an extended period of time. In other words, you want some sort of movie. Confocals can have slow frame rates for whole 2D images, and you don't want to miss calcium rises associated with action potentials. A frame rate on the order of 100 Hz is probably good, and 10 Hz is probably slow. A line or point scan of the cells you are interested in would speed up the temporal resolution.

These are my first thoughts.

-Matthew

One possible solution is to increase the amount of memory allocated to ImageJ. You can do this by going to Edit > Options > Memory & Threads and increasing the maximum memory allocation. Another solution is to try closing other programs that are running on your computer to free up memory.

Hi,

I believe it's possible if you can code in arduino and python/matlab. I have linked a paper which maybe of some help to you.

Simultaneous analysis of social behaviors and neural responses in mice using round social arena system

Hi, it is weird that you have such an hemorrhage, have you tried going down very very slow with the lens? In my lab we were just doing that. Like going down 0.1 and waiting like 10 seconds every time. And we were targeting the LH or the VTA which is pretty far ventral and never had an issue. Maybe you could try doing that if you have not tried it yet. Also 5 mm is quite a large window on the skull, it maybe to big and that may have an influence too. Worst case maybe you could try anti inflammatory drugs, I think maybe Meloxicam is useful and it works as painkiller too

Dear Orla,

I have tried Bioptech FCS2 and FCS3 microenvironmental controller with a covered bath in which you can set the temperature and feed the cells via a single inlet and a single outlet. However, it is important to check that the chamber adapter/holder will fit into your microscope's stage?

best wishes,

Refik

Ciarán McDonnell I actually had better luck when adding the dye directly to the growth media in which they had been sitting Overnight. For some reason they were much less disturbed. also, Calbryte is incredibly bright so I do just fine without washing.

at the end of the day, a new batch of cells and dye fixed the problem for whatever reason. Now everything works fine and the cells dont mind the dye at all. Very strange, but im not complaining!

Hi,

I am not sure if you tried that, but what we did was 1. create a "well" using dental cement around our window 2. on the edges of the dental cement well we applied a thin layer of vaseline to keep the water within the well. Just a tip: you should clean the objective between mice because vaseline might stay on the objective.

Good luck!

Hi there,

maybe you already solved your problem. In case you did not, here is my thought. I did some depolarization experiments myself and encountered the same problem (rat hippocampal neurons). Later, I found this publication addressing the high cysteine concentration of Neurobasal formulation leading to excitatory death of neurons:

So what I did in my experiments is to split the supernatant of the cells and use 1 half spiked up with my desired final concentration of KCL and the second half to replace the depolarization solution after the intended period of "stimulation". In my case this did the trick! Also, for any refeeding or change of medium I only used the first 10 DiV to do so with Neurobasal medium. I hope this might be helpful

Since you're measuring in epithelial cells, I assume you do not need to measure fast dynamics with your Ca2+ indicator. Therefore, I'd recommend using a high affinity GECI. This would include the "s" (slow) forms of GCaMP, which tend to have around a 1 second decay time (related to the rate of unbinding Ca2+). GCaMPs are also the best characterized. There are new and improved GCaMP versions out there that tend to be sensitive and fast (8, etc) but 6 is tried and true and gets the job done. Note this would occupy your green channel, so select other fluorescent makers wisely.

Depend on your experimental preparation, mine was the in vitro chick retina and magnesium needed to stabilize excitability. In this preparation, phosphate buffer was deadly, by contrast, TRIS was OK. Conclusion, know your preparation, read about its developpment.

I did use OGB BAPTA II already (octapotassium salt). However, I performed patch-clamp experiments (not sure if that's what you're looking for), so I loaded the cell by adding 50µM OGB in my internal solution. Worked like a charm!

Hi Frank,

By sure placing electrodes closer will help.

Also try insulating till the region which is closer to the tissue, this will concentrated the current flow.

Also consider other conductive media that may be into the bath which may form a facilitated path for the current thus impeding current floes through tissue. Leess bath volume will help.

Consider excitability issues. You may depo- or hiperpolaryze the cells varying the K extra concentration.

Probably using random noise added to the square waave pulse may contribute to activate the cells.

Best Regards,

Enrique Soto

Hello Dr Mridha,

I am really glad that the link was useful to you. I hope the will provide the results you hope for.

Best regards,

MTauro

Hello Francesca!

How to visualize the activity profile: After you open the file you can draw the oval ROIs and to confirm the ROI selection you just simply press "T" on your keyboard. Then if you go in Image > Stack > Plot Z-Axis Profile the activity plot will appear. You can select Live or not. I usually select Live because as you select new ROI it will show you the activity profile. If you like it just press T otherwise a simple click anywhere on the image will make the ROI disappear.

From the same plot you can also export the number values of the intensity profile in a separate file which later you can open with Excel or other software in order to perform all the type of analysis you need.

I use Fluo4, Fluo8 Cal-520 and Cal-590.

To be honest between Fluo4 and Cal520 I prefer Cal-520. It is way better. Lower background fluorescence after labeling and higher signal to noise ratio. I do dye injection into the tissue.

I'm not sure if I answered to your questions because I'm not sure what type of experiments you have but if you are interested to visualize the activity profile of the cells over time you can follow the steps I told you above. Good luck with your experiment and let me know if you need further help! If I know I'll be happy to help you. :)

Ciao!

Hi Sayeman,

Try stimulating with high potasium solution (ej 30 mM) they must depolarize and show a fluorescence response otherwise your system is the problem.

Best ragrds

Enrique

I’m not sure what you mean by “ratio kinetics”, so I can’t give a definite answer. To me, kinetics would refer to the speed of change in fluorescence. For that, actually, fura-2 isn’t a great dye, because it is intrinsically slow. Check out papers by Wade Regehr from the 90’s on this. If you want to study kinetics, you need fast, low-affinity dyes, like fluo-4, and you need a fast detector too, such as a photodiode or photomultiplier tube. Most cameras are too slow. It will be hard to detect a change in Ca channel kinetics even with the best system. You’ll want some solid positive controls (e.g. applying EGTA to accelerate the kinetics).

Ratio also may mean different things, especially with fura-2. If you are calculating the ratio of fluorescence at 380 nm emission compared to 340 or 360, that is telling you about the absolute concentration of Ca. You should read Grynkiewicz and Tsien about how to translate the 380/340 ratio to concentration. Additional measurements are needed. The bottom line would be, if the ratio changes with your drug, then Ca concentration is changing.

Another ratio associated with fura-2 is when you stimulate twice, and compare the amplitudes of the two fluorescence peaks. Because fura-2 has a high affinity, a ratio close to 1 would indicate the Ca concentration is quite low, in the low 10’s of nM. If the ratio is low (i.e. the dye is saturated), then the concentration of Ca is high, in the 100’s of nM to uM.

If you can give more details about your prep and measurement system, I might be able to be more specific.

-Matthew

just in case somebody reads this again. Now AAV-CAG-SomaGCaMP6f2 that is only expressing in the soma is available. That is probably the best solution for miniscope imaging:

Hi Rabeah Al-Temaimi

Thank you very much for your response! So just to clarify, I was only able to calculate F0 for cells that were acutely treated with KCl (which is our positive control) 1 min after the imaging was started. So I used the mean intensity before 1 min as F0 for these KCl-treated cells only. For the rest of the conditions (cytokine, cytokine+inhibitor, and vehicle), the cells were pre-treated with these agents 24 hours before the imaging, so there wasn't any acute addition of any agents during the imaging. I also do not have any completely untreated cells to use as F0, as you've suggested, so I am not sure what would I use as F0 (if anything) for my 24 hrs pre-treated cells? But your suggestion on using the vehicle for normalisation of cytokine treatment makes a lot of sense and I will discuss it with my supervisor, so thank you so much for it, it is very helpful!

I recommend looking into red indicators from Robert Campbell's lab (University of Alberta and University of Tokyo). See http://campbellweb.chem.ualberta.ca/publications/. Best, S

Here is the link for new miniscope version: https://open-ephys.org/miniscope-v4/miniscope-v4

Sara, the first question is when you say analyze calcium data using pluripotent cells, what exactly is your experiment? Do you expect the calcium levels to change after you stimulate the cells with an agonist/antagonist?

To put it simply, you load your cells on a coverslip, treat with Fura2, put the coverslip on an imaging chamber, and hit the cells with alternating wavelengths of 340nm and 380nm. Fura2 is a ratiometric dye, meaning the same dye shows different absorption maxima depending on if it bound to calcium or not. Free Fura2 shows a maximum absorption at 380nm compared to bound Fura2 which shows at 340nm. You plot a graph with Time on X axis and ratio of Fura2 RFU at 340nm/RFU at 380nm. Based on the treatment, if your treatment affects calcium in the cells, you should see a change in the ratio of Fura2 after the treatment. Hope this helps. Feel free to ask any questions you have.

Best,

Goutham

Hello,

You can omit step 2, and directly load your cells with fluo4-am by incubating them in DMEM.

Be careful to use PBS with calcium, as even brief moments without calcium can affect cell adherence.

Finally, have you tried to change your culture plates and use ones from another company?

Good luck

I have actually found yesterday GCalcium, an R package for this purpose.

Use ImageJ frame arithmetics, subtract background etc, ImageJ is free . kmake line or retangular profiles you will have a pixel number related to the brightness, not the amout of free calcium directly

Addition of gabazine removes this inhibition and allows the excitatory network activity to return.

What's the field of view looking like? It would be encouraging if you see at least some blood vessels. It is possible that you have a lot of scar tissue from the lens implant, or that your area is damaged. I suggest you have a look at the hystology of your animals, you could share the images if you like.

Alternatively, you can use Microfluidics systems to immobilize the whole larva for CNS imaging, while the larva is fully intact.

Dear Julius,

You may already have solved your problem, but if not I thought I'd send a few humble thoughts. Since you've tried new reagents and cells, you have ruled out some of the obvious issues such as the ATP going off or homologous/heterologous desensitisation. However, did you try a new batch of Fluo-4AM? We've had issues where the calcium indicators retain their ability to fluoresce but become less and less responsive to calcium over a few days. This happened even without freeze-thaw cycles (we make a fresh aliquot of calcium indicator for every coverslip of cells that we use, so we don't freeze-thaw). Responses to ionomycin (if it's fresh) are normally pretty reliable in our hands and so can give you an idea of how well the calcium indicator is working.

Your cells do appear to be healthy and nicely fluorescent, but the responses do seem a bit modest (although what you record on the screen will depend on the settings for your imaging system).

One comment- at the start of two of your plots the fluorescence was declining in all the ROIs recorded. Was the decline in the fluorescence due to an experimental manipulation you did just before starting the recording? If not, the drop in fluorescence would be something I'd personally need to understand. I always like to start experiments with a perfectly level baseline value for a few minutes (which is straightforward in most cells if there isn't background spontaneous calcium signalling going on). If the decline in fluorescence wasn't due to an experimental manipulation then the Fluo4 fluorescence is being lost somehow. Are you using laser light to illuminate your cells on a confocal microscope, or are you using a wide-field system? Just wondering if there is some photobleaching occurring (can happen easily with laser illumination). How long do you leave your cells after loading with Fluo4 to allow for full de-esterification of the indicator?

Apologies if this is of little use. It's sometimes hard to diagnose issues from afar.

Best of success with your experiments.

Martin

Hi Daniel,

You may have all the advice you need, but just in case here's my response to your question.

We have done quite a few long term recordings. They are not simple for some of the cell types that we study because of cell movement, which shouldn't be too much of an issue with isolated cardiomyocytes (although the cells will contract of course). The issues you may face during long term imaging include calcium indicator extrusion, indicator bleaching and photodamage. The photodamage may be caused by the illumination you use, or by ROS production via illumination of a calcium indicator within the cells. There are ways around most of these issues, but sometimes they don't work so well.

Using a ratiometric calcium indicator (such as Fura2) is ideal as a way of reducing the impact of indicator extrusion (which can be curtailed using sulphinpyrazone and other compounds) and indicator bleaching. It's not perfect though, especially if you get to minimal calcium indicator levels when the background pixel intensity and noise can become increasingly significant contributors to your recordings. To maintain calcium indicator concentration inside cells, you can do an acute loading with the acetoxymethyl ester, as Stephen said above, and then maintain a tiny concentration, say 0.1 micromolar, in your bathing solutions during your experiment. This will keep Fura2 being loaded into the cells to replace what is lost. Problem is that it could be expensive to add the Fura2-AM to your solutions. Moreover, if you lose Fura2 fluorescence due to photobleaching and have a constant reloading to compensate, the cells will have increasing amounts of a calcium buffer inside them (even if the photobleached indicator is not fluorescent).

Another issue with Fura2 is that it's not the brightest indicator with respect to fold change of the signal from Ca2+-free to Ca2+-bound. Single wavelength indicators, such as Fluo8 and Cal520, have better dynamic ranges. Also, with Fura2 you need to illuminate sequentially at 340 nm and then 380 nm to get the ratiometric advantage of the indicator. There are some fast-switching LED systems that will do millisecond 340/380 illumination (we have a CoolLED system that is very fast; microsecond switching), but most lamp and filter-based imaging systems will not be that fast.

The production of ROS and photodamage can be control for using ROS scavengers such as Trolox/vitamin E, or reducing illumination intensity/duration, but again it's not a perfect solution. It is possible to kill cells with long term illumination, as in photodynamic therapy.

Since you are interested in kinetics of calcium signals over a long period of time, I am not sure that I would use Fura2 or have a continuous recording. My approach would be to do rapid imaging of a single wavelength indicator at regular intervals (say 10 second bursts every few minutes). The calcium signals in myocytes are rapid, as you know, so continuous fast imaging for 2 hours is not that feasible. If you do your imaging using rapid imaging in bursts, you can preserve the indicator fluorescence and cell viability, and get your kinetic data at critical time points. We've used this burst 'off and on' imaging approach many times.

There is a lot of calcium expertise around you in London, but if we can help any further please get in touch.

Best of success,

Martin

It depends on you research purposes. If you are going to analyze any kind of cell or tissue dynamic then frame rate may be relevant. Otherwise not at all. . Pixel size is always relevant because it defines the resolution, but lower pixel size always means lower light sensitivity. Thus there is a treadoff between pixel size and sensitivity (light gain). Higher pixel number is desirable means larger field of view. Also lower noise is desirable in any case. And well dept is relevant because it allows to determine smaller differences in light level in each pixel. Larger is convenient at the expense probably of processing velocity or at higher computer requirements. Finally is money.....

Sorry for the delay and thank you for this helpful input about stability!

Hi Tushar,

Unless your experimental settings allow you to differentiate between the influx of calcium from the plasma membrane and the release of calcium from internal stores, your overall calcium signal will be a mix of both further affected by calcium extrusion and repumping. Therefore, a good start would be to look at the "area under the curve" which gives you an estimate of the overall calcium load.

You can of cause further dissect the various components of your calcium signal by measuring rise time, peak amplitude, decay time, half width, etc... again keeping in mind that all these parameters are affected by a mix of several phenomena occurring concomitantly.

you will need to ensure that the excitation spectra of both the indicator (voltage or calcium) and the opsin are sufficiently separate so that you don't have any crosstalk, i.e. the imaging wavelength exciting the opsin.

Our lab found a solution using Chronos for the opsin and CaSiR-1 for the calcium indicator

Hello

Kindly look at links in below

https://www.biorxiv.org/content/10.1101/308486v1

Good Luck

Dear Kwanghoon,

I also can recommend the power meters and sensors recommended by Roni Hogri and Niraj S. Desai above (I find the S130C slim sensor quite useful).

However, these devices measure *power*, not intensity. In order to measure intensity with these, you need to know the total illuminated area on the sensor. That you can achieve in a number of ways, depending on what type of light source you are using.

1. If you have a uniform beam that has a larger diameter then the sensor aperture (9.5 mm diameter for the S130C), then you can estimate intensity as total power divided by sensor area (i.e. pi*r^2). Caveat: if you beam is not uniform, this estimate will be inaccurate.

2. If you have a uniform beam that is several mm in diameter, but difficult to measure the diameter exactly, you can place a smaller aperture onto the sensor, to limit how much light reaches the sensor. For example, if you drill a 3 mm diameter hole into piece of black material and place this over the sensor, and your beam is > 3 mm in diameter, then this would allow you to calculate the intensity as described in (1) above.

3. If you are illuminating a sample through a microscope objective, then these methods may not work well (especially if using a high magnification objective that typically generates a spot smaller than 1 mm). In that case, you would need to calculate (based on objective properties, wavelength, etc) the effective beam waist diameter at the focal plane. Use this together with the total power measured to obtain intensity.

4. If you are using a fiber to deliver the light, then all you need to know is the fiber core diameter (e.g. 0.2 mm) and the total power. As others have described, you just need to make sure that all of the light out of the fiber reaches the sensor, to get a measure of total power. Be careful because the fiber tip can scratch the sensor surface if they come in contact. In this case, you are estimating the intensity *at the tip* of the fiber. The intensity at the sample will most likely be much lower and generally quite difficult to estimate. But see:

That website is currently unavailable, so here's a snapshot from 6/2018:

For consistency, if you are doing in vivo experiments with an implantable fiber, I recommend measuring the total transmitted power *prior* to implanting the fiber "cannula", as this can vary from implant to implant (and hence animal to animal). You can then adjust the command power to achieve the same power delivered to the brain in each animal.

Hi Prudhvi,

I didn't check this litterature for a while, I can remember only this article : http://www.jneurosci.org/content/17/15/5936

For the dye, I don't know this one, I used its AM homolog (OGB1-AM) wich was good but not that good compared to Calbryte.

Best

Lora

Hi Silke,

From your description I find it unlikely that the problem is in your optical system, since the raw values from your LED are many many times higher than that of your arc lamp (unless there was a change in the camera exposure, but I will assume there wasn't).

One issue of using LEDs for ratiometric assays is that there is no guarantee that your different LEDs will have the same output power. There is a chance that your 380nm LED is simply a lot stronger than your 340nm, since it is a significantly longer wavelength (so that you will have a lot more photons for the same output power with your 380nm). It might be interesting to check the output power and calculate the luminace of each LED to see if your raw values make sense.

Alternatively, you might consider switching over to a non-ratiometric method. I don't really know for synthetic dyes, but for genetically encoded calcium indicators, such as GCaMP, non-ratiometric measurement is pretty much the standard. You would need to change your LEDs to match another wavelength, but it might be worth a shot.

Hope this helps and good luck!

Hi Steve,

You wrote "initially dim neurons might weigh in more heavily". This suggests that you estimate "F0" (or what you call the background, F) from an *initial* baseline period? If so, I think you will have better results by doing what many other in vivo GCaMP experimentalists do: estimate F0 (aka baseline) over time, during the entire recording, for each neuron, using one of several common approaches. I can't tell you which approach is best, but if you dig into the methods of any recent in vivo GCaMP paper, you'll see that while the details vary from lab to lab, there are probably just 3-4 fairly common fundamental approaches.

I would say that generally one may be faced with slow-timescale drifts in baseline (which could be due to bleaching, although with GCaMP this is typically not a problem), and on a more rapid timescale, the challenge is to identify event-free recording periods wherein one can measure the baseline.

One approach is to look at the *distribution* of the signal in e.g. a 30 second window, and identify the baseline as some low percentile; or some other feature of the distribution.

Another approach is to identify Ca events, and then average across all samples outside of these events to obtain the baseline. Of course this begs the question, how do you identify events without knowing the baseline? So typically this is an iterative approach, where you start with some estimate of the baseline and iterate until reaching a suitable criterion. But there are also papers where event-free periods are manually identified, and the baseline is measured from these.

David Tank (see Dombeck papers, ca. 2009) introduced an approach that further classifies Ca events to detect false positives, based on the observation that negative events are false, and therefore the distribution of negative event amplitudes and durations can be used to predict the corresponding distribution of positive events. Essentially this allows you to assign significance values to positive events based on their duration and amplitude.

Z-score can be a useful tool as well.

Most of these approaches assume that you have a "baseline", that is to say, that the neuron is mostly quiet, and only has Ca events infrequently. This is true for almost all excitatory neurons. However, certain types of interneurons, for example, can have very high firing rates in vivo - combined with the kinetics of the Ca sensor, this can result in a trace that has no obvious baseline. If you nevertheless must use GECIs to study neurons with high firing rates in vivo, other approaches are used - I'm not particularly familiar, but a simple approach is to look at deviations around a moving-window average.

Good point Lora.

Sorry I do not have nore comment that this. My research follow the way of a Plan Electrophysiologist. I am not good enough in Organic Chemistry. However, just I am studying Synapse on Neural Network System. I am very passionate into understand, in detail, who electricity (electrones) change into chemicals from the neuron transporter to the neuron receptor.

Let us continue our conversations. We PhD students have to support each other in the usually funny way. That is life, isn't it?

Cheer,

ISA STAR-C UNI PERU

Hi Yiannis,

Looks like you get your own rig at the IGBMC, nice !

Well, purkinje are difficult to load for many reasons: size, ensheatment by glia and the presence of the MDR channel. Forget this way....

Best

Philippe

Thanks for your answer Jason!

I'm afraid I don't fully comprehend it. You mean the bulk of Ca within the animal is sequestered? I'm wondering if the 10mM Ca in seawater would have the dye chelated before it even enters the cell...?

With diluting, I'm quickly losing fluorescence. I'm already using the minimum (1mM).

Most probably also transiente oerfusion with high potassium (eg. 20 mM) solution may stimulate them by brief depolarization.

Hello, Preparative HPLC with reverse phase column or size exclusion chromatography can solve your problem.

Thank you William for your answer!

Ok thank you very much !

I'm looking for something in the cortex (S1BF or some other area) but this one helps too!

Thanks Debanjan.

Hi,

I'm using a Leica SP5 confocal for the imaging, and i use the generated TIFs and make dF over F stacks using ImageJ.

As the heart is contracting, i know for sure that there are Ca waves, i just guess the dye loading isn't good enough. how do you load the dye? perfusion of the heart before slicing, or direct culture of the slice with the dye?

I'm the first one in my lab to do this experiment. We do a lot of calcium imaging for cells, however, and never had a problem to visualize calcium waves.

Thanks

Hemi

Dear Tushar,

I share Madders opinion. It would certainly be convenient to determine whether or not your data can be fitted by more than one exponential instead of forcing fitness to a single one.

Hello Maria,

Do you have any special need for these calcium imaging experiments? I do confocal or Bi-photon imaging on a daily basis, and all the rigs I could see used standards electrophysiology perfusion systems. They are mainly DIY, with 1 or 2 peristaltic pumps and a manual flow regulator. It is really cheap and easy to do with Tygon tubes and some fittings. You can add a temperature controller of course.

The LEGO device Jonas mentionned is mainly for cell cultures, I am not certain it could be suitable for thicker tissue (like slices) where you need a higher flow rate. And it is perfectly suited for complex sequences of rinsing / Solution change, etc... but might be overkill if you only want to record your cells in standard ACSF with only 1 or 2 solution change.

Best,

Sylvain

There are some chemicals inhibitors (e.g., carbenoxolone, heptanol, or 18alpha glycyrrhetinic acid) of questionable specificity that can be used for short term inhibition of gap junctions. There are peptide based inhibitors as well which improve specificity but require large amounts to be effective ( ). Your best bet for specificity is genetic...either Gja1 siRNA knockdown or the use of cells from floxed Gja1 mice and Cre-Expression will be most convincing, assuming you get sufficient knockdown efficiency.

Hello Yang, Calcein AM does not bind Ca2+. Instead, it is converted by cytosolic esterases into green fluorescent calcein. The latter is now rarely used as a Ca2+ indicator because its fluorescence is directly sensitive to these ions only at strongly alkaline pH.

In the paper below the authors studied programmed cell death modulated by Ca2+ influx with the help of Calcein-AM.

For test purposes I would suggest trying other indicators as well. OGB-1 AM tends to behave differently than Fluo-4 AM and uses the same excitation wavelength.

Part of the behavior of AM esters when loading cells can be explained by partial charge distribution in the molecules. Thus, indicators with a striking difference in partial charge distribution will load different cell types with different efficacy.

Hey Pablo,

thank you very much for your answer! I think the buffer I use is the same, at least HEPES for my measurements and I think the problem is more about the compunds I use, et-1 and LPA. Nevertheless I figured out to handle compounds, but it was a difficulty to find out. Anyway thank you very much again and all the best from Munich!

Thanks again for the help. I'm posting this for anyone else who may have a similar issue. The issue for me was resolved for me by adding more buffer to the dish I was imaging. I was using about 1 mL in 2 mL glass dish (I wanted a lower volume to have quicker buffer transfer). I was able to observe the increase disappear while imaging after adding more buffer. I wonder if surface tension was aggravating the HEK 293 cells in some way, just a guess. The CCD did also have a minor affect so I imaged without the excitation source to prime the camera as William suggested .

@Matheus De Castro Fonseca @Florian Hetsch 

I am using HBSS for my dye loading. Post-dye loading, I prepare HBSS, where I increase the KCL concentration and decrease NaCl by the same factor. 

While imaging, I replace the HBSS medium and take basal recording (no media) and then after few seconds add high-KCL buffer. However, the cells don't look healthy and it takes >100s to reach baseline. 

With regards to movement, there is nothing you can do. You are putting a volume of non-compressible fluid into the slice. Things MUST move. You can change the nature of the movement by changing the size of the pipette (large pipettes will give you less movement over a larger volume, smaller pipettes will give you more movement over a smaller volume).

In terms of getting an instant calcium transient, I wonder what are you dissolving the drug in? If it is just plain aCSF (bicarb based) it's pH will go very strange in the pipette. Try a HEPES based solution.

When you put the puff pipette into the slice, it will get blocked. You're going to have to clear it at least once. If you don't, you'll have to use large pressures to clear the pipette, which will excacerbate your movement problem, which can cause calcium transients by themselves. If you really can't afford to get any drug on the slice, then you're going to have to "tip dip" the pipette in non-drug aCSF. This will mean the tip is filled with a drug free solution. You will then need to puff several times to get rid of that.

If none of these options are plausible, then I hope your drug can be given an electrical charge: try iontophoresis.

Dear Geoffrey,

There are no reports suggesting acute toxic effects of Fluo-4 on cells. If you are observing deterioration of the cells only after including Fluo-4 in your patch pipette solution, then the most obvious thing to do will be to check the pH and osmolarity of the final pipette solution you are using. Also since you are directly loading the cells with your patch pipette, you may need to consider reduction in the concentration of Fluo-4 you are using.

Best wishes,

Refik

To me, it looks like you filled the ImageJ frame properties correctly, and the ImageJ graph duration (x axis) seems to correspond well with the 1545 ms frame duration in your meta data.

I am not sure what you specifically want to analyse?

Do you need to setup a fully automated procedure for 100s or 1000s of experiments, or can you get away with more manual analyses and aligment of data?

What are you trying to extract from the data - are you interested in rise time, decay kinetics and/or amplitude? 

In general, I don't agree at all that "nobody cares about the precise length of the line or the LinesPerSecond". If you are looking at subtle differences between groups, then this could have a large impact on your results and distort your analyses. This will of course depend on the expected, and observed, differences between groups.

We have optimized protocol for dissociation of adult hippocampal neurons for Ca2+ recording. Please see attached.

For hypothalamic neurons, the choice of enzyme and final plating medium will be critical. See: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2578824/

Hope this helps. Good Luck!

Thanks Dr. Kulchitsky. It is very helpful article

Best regards

Hariom

I am glad to hear that :)  

Chung Sub 

Min Li,

If you use neonatal cardiomyocytes collagen would be fine and if you use adult cardiomyocytes, laminin would be best option. Becasue of the hydrophobicity of the glass water/buffer would not form a thin layer. So repeated pippeting until it form a layer on glass would work.

Dear Bapon,

There are several possibilities why you are not seeing any ATP responses. Your ATP is infected by bacteria and chewed up. Or your transfection did not work and your cells are not expressing any ATP receptors. Or your cells are are depolarised and resting membrane potential close 0 mV and therefore the cells are not responding to ATP. I have done expression in oocytes before and studied ATP-gated ion channels. See my publications listed below.

best wishes, Refik

thank you..:)

Nicola, there are a number of ways in which you can gain good functional insight into synapse properties without using electrophysiology. As you suggest in your question, fluorescence imaging would be the way to go about this. I personally would prefer using optical methods to electrophysiology when monitoring synapse function. I suppose your are not yet set on what exactly you would like to monitor, so I will just provide a few general ideas.

 In cell culture the use of a good wide-field epifluorescence system, or a (spinning disk) confocal will allow large numbers of cells and synapses to be assessed in a relatively short time frame. There are numerous probes available, that provide access to different synaptic properties -- it is simply a matter of selecting which parameters you need or would would like to quantify:

Fusion proteins targeting pH-sensitive GFP-variants to the inside of synaptic vesicles can be used to monitor fusion events. Such probes are dim in the acidic environment of the vesicle, but light up once exposed to the neutral pH of the extracellular space, thus lighting up on vesicle fusion. SypHy or Synapto-pHlorin and it's variants are the most commonly used probes for this.

Calcium indicators can be used to monitor both pre- and postsynaptic calcium influx. Loading cells with AM-esters of small molecule calcium indicators such as Fluo-4 or Fura-2 is one option. But a number of genetically encoded probes are also available. GCaMP6f (the fast variant is better suited for imaging synaptic events) is the most commonly used probe for this. Fusing GCaMP to Synaptophysin results in the indicator only being present in the presynaptic boutons, increasing the signal to noise of the measurements and allowing for clearer interpretation of the acquired signals (SyGCaMP, Lagnado lab). In addition, red genetically encoded calcium indicators are also available, such as R-CaMP2 (from the Bito lab) which should make it possible to image pre- and postsynaptic activity at the same time.

It is also possible to image the release of neurotransmitters. Glutamate indicators, such as iGluSnFR (Looger lab at Janelia) are the tools used for this. These signals have a much higher temporal resolution than imaging pH or calcium transients, which might make them a better match for your experiments.

Using fusion proteins it is also possible to monitor changes in receptor numbers at a given synapse. This can be achieved by assessing brightness changes (careful controls are needed here) or ratioing brightness of different color proteins fused to different proteins. For example, AMPA-receptor internalization can be monitored by fusing a pH-sensitive GFP-variant to the extracellular part of the receptor, leading to a decrease in fluorescence on internalization, similar to the Synapto-pHlorin approach mentioned above.

To get deeper into the biophysics, fluorescence recovery after photobleaching (FRAP) could be used to monitor vesicle or protein mobility within a synapse. A bit over the top, cellular fluorescence correlation spectroscopy (FCS, see the work of Petra Schwille among others) can be used to monitor the concentration and diffusion rates of synaptic proteins.

I hope I have been able to provide you with a few useful pointers, while not going into too much detail. As you can see, the sky (or your imagination) is the limit.

Good luck with your experiments!

In addition to Norbert's helpful suggestions... this can happen if compounds affect Fura2 directly. Caffeine has such an effect (Biochemical Journal 329, 349-57). One way of checking this is to deplete all intracellular calcium stores (e.g. with application of ionomycin in calcium-free medium) to prevent any calcium flux in response to your agonist. Following that treatment, if your agonist still changes the 340/380 ratio then it could be a direct effect on the indicator. Can you switch to a different calcium indicator (e.g. Fluo-8, or a ratiometric pair of Fluo-8 and FuraRed)? That might alleviate the issue.

Can you show us the 340 nm and 380 nm traces rather than just the ratio? If it's a genuine calcium change there should be corresponding reciprocal changes in the 340 and 380 traces.

Best wishes,

Martin

See our paper  

Fujiwara, T., Kazawa, T., Haupt, S. S., & Kanzaki, R. (2009). Ca2+ imaging of identifiable neurons labeled by electroporation in insect brains. Neuroreport, 20(12), 1061-1065.

We used Ca-green-dextran 3000 not fluo3/4.  But if you can insert glass electrode to some specific region (MB?)  adequatel,  it  may be possible to stain some specific type neurons.

Dear Roberto,

Classically(patch clamp and calcium imaging) we used platinum harp which is heavy enough to stick the slice on the bottom of the chamber, but which is not ionizable. THere are many commercially avaible harps that perfectly fit in the recording chamber. However, you can also make it yourself.

Pascal

5 uM loading for 30 minutes at 37 C. Wash twice.

What lasers/detectors are you using? Have you used a positive control?