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Calcium Imaging - Science method

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Hi,
I am currently working on optimizing the protocol in my lab for 2P imaging using a water dipping objective. I'm using mice with implanted GRIN lenses that are 0.6x7.3mm, imaging amygdala neurons (injected at approx -4.7mm depth from bregma). I'm having trouble maintaining the meniscus between the objective and the lens. Are there other compounds that people use or any methods to maintain the meniscus that I may be unaware of? Thanks for any advice!
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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!
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Dear colleagues, I’ve started working with cultured primary neurons and came across a problem.
I need to depolarize neurons for different time intervals (up to 1 hour) and then use them for an assay 2 days after. The problem is that most of them are dying after depolarization.
I culture neurons in complete neurobasal medium (Neurobasal + 2% B27 supplement + 1%Glutamax +1% penstrep) with 1/3 media change every 3 days. I depolarize them on DIV11-14 by swapping media on Tyrode’s solution (45mM KCl) for up to an hour. Then I wash the cells with Tyrode’s solution (5mM KCl) twice and swap the collected media back.
45mM K+ Tyrode’s: 100 mM* NaCl; 45 mM KCl; 1 mM MgCl2; 1.8 mM CaCl2; 1.04 mM Na2HPO4; 26.2 mM NaHCO3; 10.9 mM HEPES; 10 mM D-glucose
* NaCl is used to adjust osmolarity of the solution, so concentration varies.
Since in the current setup I depolarize cells in 5%CO2 incubator I used buffering formula of Neurobasal media. I adjust the pH to Neurobasal’s pH=7.7 and checked that in CO2 incubator it equilibrates to pH=7.4. And I adjust solution’s osmolarity to match the current neuron’s medium too.
Also, I depolarized neurons in live cell imaging using GCaMP6s to monitor calcium elevation and after minutes I can see that some neurites are destroyed (Fig.1, attached) and after 0.5-1 hour cells don’t look good and most of them die afterwards (Fig.2).
At the same time, I keep seeing papers with no explicit details on solution and osmolarity where cultured primary neurons are stimulated with KCl for hours. For example, here 6 hours of 55mM KCl (https://www.nature.com/articles/nature09033).
I guess there are a lot of people routinely working with primary neuronal culture. Could you please help me, what am I missing?
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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
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I would like to express a GECI in primary epithelial cells. I Would like to get some suggestions for to chose the best GECI reported. Are there some GECIs reported such as GCaMP5G, GCaMP6s, GECOs, but I don't know what is better than other.
Thanks in advance.
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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.
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Hi there!
I'm trying to perform an experiment to prove one specific protein interacts and activates NMDAr. For this purpose i'm trying to use Fura-2AM dye to see changes in Ca2+ levels inside the neurons and two photon microscopy. As a positive control, I used different concentration of NMDA (from 0.1 uM to 100 uM) which I applied to the medium with neurons. Surprisingly (?), i see little activity from the neurons.
I noticed that usually people use magnesium free medium while performing this Ca2+ measurements for NMDAr activity. I suppose the reason being Mg ions block the channel and Ca2+ cant go thru. I'm just curious if 1) Mg presence in the medium could completely explain almost no activity from my neurons in regards to Fura 2AM intensities and
2) what is the relevance of such an experiment when my other experiments were done in normal neuronal medium (with Mg ions)? Can I claim effects of the protein X is due to NMDAr binding and activating when I can't get any activation signal unless I use magnesium free medium?
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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.
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Good afternoon, I am trying to do calcium imaging of MDA-MB-231, U87, and CT26 cells using Cal-520 AM dye which is by all accounts nontoxic to cells, and I have used it for years with no issues...
However, recently, I've noticed that many of the cells become rounded and appear unhealthy after 1hr incubation at 37c. They do not appear to recover after the wash and RT incubation step either. This only began in recent months, but nothing obvious has changed about my protocol/cell types, and I just cant figure out what is causing it...has anyone ever seen such dyes causing cells to round up like this?
Already tried without much success:
- switching dye batches (old to new)
- Incubating in complete growth media/KRB/Hanks + 20mM HEPES
- adjusting concentration between 1, 2.5, 5uM which is the recommended range. I cant see a difference in degree of rounding, but the signal is too weak to use below 1uM...
- Different cell types
- varying cell density
- shorter incubation times. This appears to happen by about 30mins or so, but I haven't yet done a proper time course to know for sure.
Can anyone offer any suggestions?
Thank you in advance
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Thank you for the helpful suggestions! I doubt that it’s a batch issue because im having the same problem with brand new aliquots (stored at -20) as with the previous batch which was almost a year old. I never considered the DMSO before because it supposedly doesn't go bad, but it’s ~1.5yrs old and was used for both dyes so maybe i’ll order a fresh DMSO batch…
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Dear all,
We are starting a project that involves calcium imaging from activated T cells; in the literature people often use ratiometric dyes but I prefer to use OGB or calcium green AM
Does anyone has experience in loading theses cells with these dyes? can anyone reference a previously tested protocol? Many thanks
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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!
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Hi, we would like to analyze time sequences images of calcium imaging assay performed with Fluo4-AM in imagej. Is there any plugin that we could install to do this? Do you have any kind of suggestion? we are interested in obtain a graph showing the activated cells over time, like a raster plot. Is there the possibility to automatically identify the intensity spots? Thank you!
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Qui puoi seguire passo passo. Here you can follow ste by step:
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I am currently using the Warner RC-49MFSH (https://www.warneronline.com/perfusion-chamber-with-field-stimulation-rc-49mfsh) which has two platinum wires placed parallel approximately 10mm away from each other. I have a Grass SD9 Stimulator connected across these electrodes, which allows me to generate pulses of varying parameters (0.2-200Hz, 0.02-20ms, 0.1-100V). ~1 mm below my electrodes I have a glass coverslip, upon which I place an isolated murine dorsal root ganglion (DRG) (total length of say 1 mm), with a nylon mesh placed on top to hold it in place. I then immerse the tissue and the two electrodes in artificial cerebrospinal fluid (aCSF). I then do calcium imaging on the neurons in the DRG.
My problem is that whereas previously the lab members were able to see a high amount of responding neurons, after we switched a strain of mouse (from Advillin promoter to Thy1), I have had very spotty results with responsiveness in the neurons, despite testing a wide range of parameters and going up to the maximum voltage allowed. I might see one or two neurons respond, as opposed to the ~100 that was found previously.
What are some factors that might affect the magnitude of electrical stimulation on tissue in this setup? I saw some other threads that suggested moving the electrodes closer together, which I plan to try very soon. I also checked the resistance across the electrodes and the aCSF bath and found it to be in the range of 100 kOhms to 1 MOhm. The other thread suggested sanding the electrodes in this case. Should I also minimize the volume level of the bath? Does the length of wire immersed in the bath affect the total current flowing through? What happens if I put the electrodes so close together that I can physically touch the DRG to the electrodes? Should I place the DRG closer to the positive terminal since the positive terminal should attract negative ions in the solution, thereby depolarizing the neurons?
I know this is a lot of questions and such a system can get complex quickly. I am still working on understanding the electrode/electrolyte interface. Thank you in advance.
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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
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I am trying to do calcium imaging from mouse adrenergic axon (NE).
I injected “AAV5-hSynapsin1-FLEX-axon-GCaMP6s” to the LC nucleus in DBH Cre and TH Cre mice, unfortunately have not seen adrenergic axon in the cortex.
Can anyone guide me to find an appropriate and efficient AAV to do calcium imaging from mouse adrenergic axon?
Is there any new AAV for Cre dependent, axon targeted GCaMP ?
Thank you
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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
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Hi everyone,
I am trying to measure calcium sparks and transients in freshly isolated rat mesenteric smooth muscle cells using Fluo-4 (5 microM). we have a Zeiss LSM 900 confocal microscope that we use to image the cells.
Unfortunately, cells don't show any change in florescence after the administration of drugs. We have tried Angiotensin-II, caffeine, Calcium Ionophore (A23187) with no luck.
Has anyone faced any similar issue like this one? Does it mean the Fluo-4 has gone bad? I dissolve Fluo-4 with DMSO right before use. Fluo-4 loading is fine, since i can see the cells in AF-488 light.
Any help will be deeply appreciated.
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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
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I am new with this technique. It seems that different cells are responding to my drug with a very different ratio kinetics in my calcium imaging experiment. do they mean anything?
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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
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Which approach is better for in vivo calcium imaging in the brain with miniscope: image in virus-injected (AAV GCAMP6) or transgenic GCAMP6f animals?
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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:
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Hello,
I am new to calcium imaging data analysis, and I need to analyse it for my master's project (I didn't actually acquire the calcium images myself). So in the experiments, the primary mouse neurons were pre-treated with one of the three possible treatments (vehicle, cytokine alone, and cytokine + its receptor inhibitor) 24 hours prior to the imaging. Then, on the next day, the neurons were loaded with Fluo-4 calcium indicator and imaged for 5 mins. KCl treatment was also used as a positive control, but unlike with other treatments, neurons were not pre-treated with KCl for 24 hours. Instead, KCl was added to the neurons 1 min after the imaging was started. I used the software for the microscope to obtain the time measurements of changes in Fluo-4 intensity separately in somas and neurites.
So for KCl, I was told to use the mean of the intensity before KCl addition (i.e. 1 min) as F0 and divide all the intensities by the corresponding F0 values (i.e. F/F0), but I am not sure what to do with the other treatments since there isn't a clear period which I can use as an F0 to normalise them. I am also not sure if I need to normalise these at all. I would also appreciate any suggestions on the type of statistical analysis that I should use to compare these treatments.
Thank you in advance for your help!
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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!
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We regularly use Gcamp6s in our lab for in vivo calcium imaging. The problem is that we have a mouse line with GFP marking a specific cell population that we want to image in vivo and hence we need a different coloured GECI.
Specifically, I want to know if the Douglas Kim lab Rcamp indicators are sensitive enough to use in 1 photon recordings. Further, how well do they compared to Gcamp's such as Gcamp6.
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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
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It is a version of Mark Schnitzer's design for a head-mounted microscope to monitor fluorescent calcium signals in freely behaving rodents, published in Nature Methods in 2011.
I am looking for general feedback on putting it together and using it as well as comparisons between Miniscope and Doric Lenses or Inscopix's much more expensive systems.
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Here is the link for new miniscope version: https://open-ephys.org/miniscope-v4/miniscope-v4
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I am new to calcium analysis and I have to analyse calcium imaging data of human induced pluripotent stem cell derived cardiomyocytes. Does anyone know any good tutorials/papers or have any guidance for beginners?
Thank you for your help.
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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
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Hi everyone,
Currently I am trying to calcium imaging using HEK293T cells,
however, I can not measure fluorescence because HEK293T cells can easily be stripped after following procedure,
I already tried to use collagen coated plate and zelatin coated plate but both did not work as well.
Could anyone teach me good method if you are familiar with that, thank you so much
My procedure is following
1, seeds HEK293T cells 4×10^4/well to 96 well plate
2, remove culturing medium and wash with PBS.
3, Add loading buffer (recording medium + Fluo4AM probe), incubate at 37℃, 1hr
4, remove loading buffer and wash with PBS
5, Add recording medium
6, Add ionomycin to induce calcium ion inside of the cells
7, measure fluorescence
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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
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All the tools I found deal with the processing from the raw data to the calcium components, but I am interested in the post-processing.
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I have actually found yesterday GCalcium, an R package for this purpose.
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Hi,
I have 2 3d tiff stacks collected by imaging along axial plane of mouse brain area in vivo from different days, before and after fluorescent viral injection.
I want to detect changes in brightness of the 2 image stacks (mostly has dendrites only). What tool must i use to do this?
I suppose background subtraction, dennoising, applying filtering and image normalization must be done prior to the above detection. The 2 tiff stacks have histograms that are not completely matched (mean, std dev, min max of image stacks are different, though not very much)
Plus, help me out here please - is the attached image noisy ? i see fluorescence images in papers have a black background and dendrites/neurons in color in foreground. For the most part, they appear clean (no background fluorescence).
Is this a problem caused by imaging settings during the experiment ?
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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
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I am trying to characterise GABA developmental switch in iPSC derived neurons using Ca2+ imaging. However, as I add GABA agonist, I observe the spontaneous activity of several neurons being abolished, and addition of gabazine in those neurons brings up the transients again.
I am only able to find fewer neurons in culture that show Ca2+ elevation upon GABA addition.
Any idea why this could be happening?
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Addition of gabazine removes this inhibition and allows the excitatory network activity to return.
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Hello,
At the moment I am trying to implement the miniature microscope (more specifically the Optogenetically Synchronized Fluorescence Microscope System - Deep Brain, from Doric Lenses) and, consequently, calcium imaging for this system.
Presently, we are facing some problems regarding cell firing visualization. Until now we haven't been able to visualize any cell firing, either in the presence or absence of tone and shock pairing, during fear conditioning.
We have infected mice amygdala with (0.5ul of) jRGECO1, an mApple based virus. In the same surgery where we introduced the virus we also implemented the canula approximately 50um above the injection site.
Did anyone face the same problems or have any suggestions regarding any possible optimizations or why we are not visualizing cell firing?
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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.
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I'm looking for a simple and quick way to immobilize Drosophila isolated CNS before Ca imaging without movement disturbance of the brain due to liquid containing stimuli application on it...
Any idea? tips?
Thanks.
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Alternatively, you can use Microfluidics systems to immobilize the whole larva for CNS imaging, while the larva is fully intact.
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Hi!
I’ve been imaging ATP-evoked Ca2+ response from skin fibroblasts. I’ve used 80uM ATP to evoke the response in 0-Ca2+ HBSS. The protocol I used worked fine at first, but when I continued the experiments after a short break, I haven’t been able to get the cells to respond to ATP. On top of the missing ATP response, cells don’t have spontaneous activity at all. When stimulating the cells with the 10uM Ionomycin in the 0-Ca2+ HBSS, the Intensity raises, but relatively little (about 2X baseline values). The cells seem to be healthy and in good condition.
I’m using Fluo-4 AM dye, and I have incubated cells for 50 min RT before imaging.
I can see change in the fluorescent intensity when changing the extracellular media from 1.8M Ca2+ containing HBSS to 0-Ca2+ HBSS, an vice versa.
I have already checked all the reagents and tried to image new cell lines, too, with similar results. The response for the bradykinin is missing, too.
Thank you for the help.
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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
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Hi,
I'd like to measure the cytoplasmic calcium transients in single cardiomyocytes upon increasing concentrations of a drug over a time of 1.5-2hrs. It should be a continuous recording throughout the whole time, so I'm worried about potential cytotoxic effects during the experiment.
I've never done any calcium imaging before and the sheer choice of different dyes and genetically encoded sensors is a bit overwhelming. What's the best choice? Does anyone have specific recommendations?
Some more details:
I'm just interested in the calcium transients, so I don't need other channels for additional fluorophores. I also don't need to know absolute calcium concentrations, I'm just interested in the kinetics.
Thanks for any advice in advance!
Daniel
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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
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What factors are MOST IMPORTANT to take into account when purchasing a camera for a light field imaging system (i.e, QE, frame rate, sensor / pixel size, overall pixel count, read noise, well depth, etc.)?
This microscope will be used for neuronal calcium imaging in small organisms (ciona, c. elegans, zebrafish).
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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.....
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Hi all,
I would like to infuse a drug (CNO) and monitor neural activity in the same brain region using calcium imaging (fiber photometry) in mice.
Ideally in the in vivo behaving mouse. But I could start with anesthesized mice.
N.B. I can inject CNO, wait 1/2h, and then run fiber photometry.
I have been using cannulas from Plastics One (Invivo1) for local CNO infusions. Typically in the lab my colleagues have been using Doric fiber photometry systems.
Did anybody ever combine the two ? I see Doric has optogenetic/drug infusion dual systems but I couldnt find fiberphotometry/drug infusion dual systems.
Or maybe I should just make it myself...?
Any input or ideas will be highly appreciated !
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Sorry for the delay and thank you for this helpful input about stability!
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I am stimulating my cultured cells and recording calcium activity as fluorescence change using live-cell calcium imaging. Though I am more concerned about the decay of the signals, for which I am calculating Tau ( the time constant of decay).
Additionally, I am interested to examine the Rise of Signal. What would be the best parameter to study a rising signal? For example, a few measures of a rising signal are Rise time, Time constant, Slope ..................etc...
P.S. Not having a mathematical background :P
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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.
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Please provide solution that is readily available. Also what is the downsides of each of them when comparing to electrophysiology?
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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
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I routinely functionally assess the iPSC derived neurons using calcium imaging with Fluo-4/AM (490nm/510nm). However, have not been successful at using those same neuronal dishes for carrying out ICC. The reason that I think could be are as follows:
  1. The calcium-sensitive dye fluo-4/AM has an emission at 510nm. As a reason why, there is a spectral overlap with my anti-bodies of interest.
  2. As I try and wash out the dye from the dishes, the neurons detach as a matter of fact. These neurons have been subjected to constant washing before and after the calcium dye-loading process. Moreover, these neurons are first stimulated with TTX following ionomycin and EGTA+TX are added to the dishes as internal controls.
Any suggestions or references or direction to a protocol would be of a great help!
Thanks!
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Hi.
I've been using LED for optogenetics and calcium imaging.
For now with the devices, I'm only able to adjust the currents or % power of the LED as out power can vary from patch cord to patch cord.
While it seems like people usually adjust the power of LED by W/mm^2, how would I be able to measure that?
What kind of meter do I need?
Please recommend me if you've been using one.
Thank you!
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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.
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Hello everyone,
I am doing single cell two-photon Calcium imaging using whole-cell patch clamp. Briefly, I am patching CA1 cells with an extracellular electrode positioned on the schaffer collaterals to stimulate the CA3-CA1 pathway. I use Fluo-5F and line scan a ROI to detect the calcium transients along the different dendritic positions to see the corresponding change in Ca fluorescence with stimulation. I see a clear corresponding change in fluorescence when I do a line scan along proximal dendrites but fail to see any signal when I move towards distal dendrites. So my questions are the following.
1) Do Ca kinetics differ from proximal to distal dendrites?
2) Is there any other dye suitable for imaging the Ca fluorescence in the distal dendrites?
Any suggestions are welcome and thanks in advance.
P.S. I also tried using Fluo-4ff but it didn't work as well.
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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
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Hello,
I have to build up a new calcium imaging system which has 340nm and 380nm LED´s (should be very new on the market). When I load my cells (e.g. mouse neurons from DRG´s or recombinant cells) with Fura2 (3µM for 45 min. incubation) I get nearly no signal. What I noticed is, that my baseline starts at a very low ratio (~0.05). I have rowdatas of 500 at 340nm wavelength, and 8000 at 380nm. Normally the baseline is between 0.6-0.9, isn´t it? I also have an older Setup with an arc lamp. There I normally have rowdatas of 150 at 340nm and 200 at 380nm. Could it be that my intensity of the 340nm is too low? Or is the 380 nm too strong?
Does someone knows what my problem can be? I´m grateful for any idea!
Thank you!
Silke
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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!
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I'm 2-photon imaging stimulation-evoked calcium responses of deep neurons using a transgenic GCaMP6s mouse line. The neurons are quiet, meaning their baseline Fluorescence (F) is rather low. The fact that they are deep also reduces the baseline (F). I notice in calculating (delta F)/F [the change in fluorescence (delta F) normalized to the background (F)] that some small responses actually give massive values. This is because they have a faint baseline (F); as the background approaches zero, (delta F)/F would go to infinity, meaning initially dim neurons might weigh in more heavily, especially if some pixel values are zero (absolute black). This doesn't seem right. Does anyone have comments on the limitations of delta F/F? Are there stress tests to determine if it is valid for certain preps? Alternative methods? Suggested reading? When we pool the responses of many neurons to make generalization about the population, which neurons should weigh in more heavily?
I might normalize the calcium response to a red protein expressed under the same promoter as my GCaMP6s. The units would be arbitrary, but it seems useful for comparisons.
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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.
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Hi everyone !
I am performing acute slice Ca-imaging and I would like to register the fluo rate increase on neurons in response to DHPG(
group I mGlu receptor agonist). Does somenone know if I can stimulate the same slice multiple times with DHPG ? Or does only the first stimulation is reliable ?
I obtain very variable responses but I don't know if it's correlated with the number of stimulations or if the responses are classically very variable in this experiment...
Thank you for your help
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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,
Victor P ROJAS Yupanqui
ISA STAR-C UNI PERU
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Hello everyone,
I am using acute cerebellar slices (200 um) to bath load them with the next generation calcium indicator Calbryte520 AM. I have used several optimizations but what I am getting is the following: A very nicely stained cerebellum with strong signal in Granule cell and Molecular layer, but my Purkinje cell layer (Purkinje cell somas) is not stained at all. Do you know why? Is it due to limited retention of the dye in these particular neurons? My literature research yield to very little information so far. Does anyone faces the same problem or managed to solve it already? Thank you in advance.
Best,
Yiannis,
PhD student at IGBMC,
Translational Medicine and
Neurogenetics.
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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
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I am trying to calcium-image marine invertebrate larvae, by incubating larvae in FSW in AM-ester dye (Fluo-4 AM).
I am wondering if dye saturation is the reason I'm not observing any signal change. How do I overcome the issue of the dye chelating free Ca2+ in FSW prior to entering target cells?
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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).
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I want to measure the intracellular calcium (using fura-2) concentration after knock down of my gene of interest using siRNA. Whatever I have gathered based on reading protocols, H9C2 cell needs to be stimulated either by caffeine or using electrodes. I want to know is it absolutely necessary to do so?
Also, since I want to measure resting intracellular calcium concentration after siRNa silencing of my gene, should i go for microscopy or plate reader based assay like FLIPR?
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Most probably also transiente oerfusion with high potassium (eg. 20 mM) solution may stimulate them by brief depolarization.
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I have cell conditioned media which contains a protein or metabolite that causes a significant calcium response in another set of cells. The first step towards identifying the factor is to split the conditioned media into fractions, however, I would need to then test each fraction separately in calcium imaging experiments to see which fractions contains the factor that I am searching for., before proceeding to proteomic analysis of the fraction of interest.
I have looked into using simple centrifuge filters, but from what I gather they are more useful for concentrating proteins rather than reliably splitting them into fractions based on kDa. would it be possible to use reverse phase HPLC for this purpose? is it possible to acquire distinct protein fractions from this method which I could then re suspend in media for calcium imaging experiments?
Any suggestions would be greatly appreciated, thanks
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Hello, Preparative HPLC with reverse phase column or size exclusion chromatography can solve your problem.
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I'd like to have some advices to perform calcium imaging in embryonic neuronal cultures.
Thanks to anybody who will help me
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Dear Greta, depending on my experiences with calcium imaging in embryonic stem cells derived from cardiomyocytes . Accordingly, you can use the recommended concentration 0.5 micromolar for about 30 mints. Regards
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Hello All,
I am trying to do calcium imaging in primary cortical neurons at long-term time points like 6 hrs after imposing treatment. Does anyone know how fast can calcium dyes stay in neurons before being transported out?
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Thank you William for your answer!
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Hello, does anyone know a way to identify oxytocin neuron over the vasopressin neuron in acute slices ? I am performing calcium imaging in the PVN of mice with Calbryte dye and recording calcium spikes but I don't know how to diffrentiate the responses from the OT neurons over the VP neurons.
As I'm working on specific KO mice, I can't use GFP-OT mice...
Thank you for your help !
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Ok thank you very much !
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Hey guys,
I am trying to find the right literature on calcium imaging studies of PV interneurons.
Could anyone point me in the right direction?
Would be even better if the studies looked at early stages of development (right from P6 onwards)
Thanks in advance!
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I'm looking for something in the cortex (S1BF or some other area) but this one helps too!
Thanks Debanjan.
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Hi all,
I'm using a vibratome to cut heart slices, and I want to do calcium imaging of the slices. I know for sure that the tissue is alive, as it is obviously contacting upon stimulation (without BDM).
However, I can't get any calcium waves from it. I stained the slices with fluo -4 or Rhode 2 (5 uM, with pluronic), and also tried to load the dye using layendorff perfused heart and then cut it to slices.
Love imaging didn't give any flurescence changes.
Will be grateful for any suggestions.
Thanks a lot
Hemi
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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
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I am stimulating my cells in culture using a variety of stimuli and checking the response as Calcium Intensity changes, using live cell calcium imaging.
The rise is instantaneous but decay seems exponential.
I am fitting this decay using exponential decay model in MATLAB and getting an R-square estimate.
What R - square value is considered a good fit ?
Should i discard the exponential decay curves, below a certain value of R-Square ?
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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.
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I want to buy a perfusion system for a confocal microscope. Would you recommend one? Mostly, I want to do calcium imaging. I would like one that you can control the flow rate. Thank you so much for the advice!
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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
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With reference to a publication on Inhibiting Cx43 Gap junctions in cardiomyocytes using chemical compounds such as Ioxynil octanoate, I would like to inhibit gap junctions and image live-cell calcium imaging to observe the effect. Can someone provide me suggestions?
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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.
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Recently, I want to detect the cardiomyocyte cell viability including live cell staining, total cell nuclei staining and apoptosis cell staining. Also, in order to show the function of cardiomyocyte, I need to get the Ca2+ flux, by Ca2+ imaging. For the live cell staining, I want to use the Calcein AM. For the Ca+ imaging, I want to use Rhod-3 Calcium Imaging Kit. Is this double staining suitable? I see the principle of Calcein AM staining and Rhod-3 Calcium are much similar. They will bond to Ca2+ and give off fluorescence signal. Could I just use the Calcein AM to take the Ca2+ imaging? I am beginner of cell staining. So I have several problems.
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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.
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Hi all,
I'm currently working on getting calcium imaging working on my rig. Currently, I'm using 50 ug alloquots from thermofisher and following their electrophysiology protocol which includes the addition of 44 uL DMSO and 9 uL pluronic acid. I then apply the Fluo-4 directly to the slices and incubate for 40 minutes. I was wondering if anyone else had experience with this, as their electrophysiology protocol is for cultured cells. Under the scope, I do have cells that are labeled, but I cannot seem to get evoked responses from stimulation or stimulation mixed with a high potassium solution. I've found a couple of JOVE videos discussing the methods, but most are using the Fura indicator.
Best,
Brandon
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Hi Brandon, have you solve the problem. We are using fluo4am as well and we met the similar problem. neuron is loaded with dye but give no response to electronic stimulation.
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Hi all,
I'm doing Calcium Imaging experiments in murine lung fibroblasts. Therefore I use several different compounds, like LPA (250µM) and Endothelin-1 (10µM). By using those compounds in a HBSS solution with Calcium I got trouble to find the right setup, solution and concentration. I tried to handle Endothelin-1 in different ways, for example on Ice between the single runs, but it's strange, because sometimes I can see a response and sometimes not. For LPA I had to use quite a high concentration to get a consistent response. So, my question is if anybody knows how to handle endothelin-1 and lpa in Calcium Imaging experiments? Any suggestions or comments can help! Thank you in advance,
Larissa
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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!
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I transfect HEK293 cells in DMEM 10% FBS (D10) with .75 ug of DNA of gCaMP6s and let them grow for 24 hrs. I then transfer the cells from D10 a commercial imaging buffer from ThermoFischer. I wait 20 minutes, I image the cells at a rate of 1 frame/second over several minutes. I see a slight increase in intensity over minutes in a majority of cells (About 5 % increase over 10 minutes on average). The cells are not flashy, it is simply a slow linear increase in brightness. Any thoughts? Thanks
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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 .
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Does one add the high-concentration of KCL (~40-50mM) solution directly into the dish as its being imaged?
Does the calcium-dye loading needed to be done in the above solution as well? 
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@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. 
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Hello everyone,
I am currently performing calcium imaging experiments in rat acute brain slices. I have been doing this type of experiment for a while now and never had any problems with bath applications of drugs. However, I am now working with drugs that I cannot apply via bath because they are very expensive. Therefore I am trying to apply drugs with a patch pipette positioned near the cells of interest.
However, I am facing wo major issues :
- the pipette gets clogged very often (unlike with patch clamp, i cannot apply a positive pressure when going into the slice because it would apply the drug too soon and desensitize my receptors). Therefore I'm not always sure I do apply the drug. To be certain of whether the drug is applied onto the slice, I added sulforhodamine 101 to the pipette to monitor drug application, but it doesn't fix the clogging issue...
- I get a lot of purely mechanical responses to the application, even with very low pressure applied . Some cells display calcium transients almost immediately after the start of the application, whereas the response I'm looking for should take longer since it involves GPCR activation).
Does anyone have advice on how to overcome these issues ? Thank you in advance !
Ivan Weinsanto
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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.
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I am loading pancreatic islet cells with ThermoFisher's Fluo-4 pentapotassium salt via my patch pipette in order to image calcium. I experience a loss or deterioration of my seal after going whole cell usually within 10 minutes (sometimes faster). Has anyone used Fluo-4 in this way and know whether it can compromise seal integrity please? Thanks in advance. 
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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
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Hi all,
I am currently doing some experiments using whole-cell electrophysiology combined with two-photon imaging to study calcium dynamics. However, when it comes to the analysis of line scans, I am unable to align/match the time points of different traces as each trace has its own/different time points (which in turn depends on the length of the line during acquisition of the trace).
Could anyone with expertise in this field help me how to analyse the traces? At the moment I am using ImageJ for doing the analysis.
Any answer would be appreciated. Thanks!
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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.
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Does anyone know of a protocol for dissociation of adult mouse brain hypothalamic neurons to be used in acute calcium recordings? We are not interested in doing cell culture, but rather using the cells acutely after dissociation.
Thanks!
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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!
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Any calcium imaging or similar techniques to measure nNOS with its agonist and inhibitors will also we appreciated. Your advice, recommendations and references will be highly appreciated.
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Thanks Dr. Kulchitsky. It is very helpful article
Best regards
Hariom
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Hi everyone, my labmate and I have been trying to clone a few shRNA constructs in the past 2 months. We encountered quite a few issues and only successfully cloned 2 out of 18 constructs so far. I would like to ask for some advice from you. Thank you so much in advanced! Below are my questions and link to brief description of our protocol + result. Please let me know if there is anything else you'd like to know. 
Best,
Huong
- Our high background indicated that the CIP didn't work very reliably and that the vectors were not fully digested by both enzymes. How to fix this? Maybe to use other phosphatase? serially digest the vectors? 
- Do you have recommendations on how to make sure CIP works properly? If not CIP, how to efficiently get rid of/reduce the background?
- What is your protocol for annealing the oligos (temperature setting, buffer, concentration to start with, etc)? 
- How effective is the phosphorylation with T4 PNK? How to tell if the reactions actually work? Maybe by running gel?
- Did you follow NEB recommendation for total DNA concentration during ligation? If not, what did you usually use? And what is the optimal ratio of vector:insert?
- Out of the 9 positive clones that we got, 3 of them have some sort of mutations. Could the bacteria be accounted for this? If so, which other competent cells are better?
- Our successful constructs are for the same domains on two proteins. Could there be a favor toward these oligos? How to enhance the success rate of the others?
Link to protocol and result is attached! Thank you all!
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I am glad to hear that :)  
Chung Sub 
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I work with neonatal rat ventricular cardiomyocytes, and I want to do calcium imaging of the same. I have been trying to attach cardiomyocytes to glass coverslips and I have tried a few substrates like laminin, fibronectin, collagen and gelatin. But the cells in the glass coverslips detach or die on the second day of the culture whereas those in dishes remain healthy. Has anyone tried attaching the cardiomyocytes to glass coverslips? Could you suggest to me a substrate?
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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.
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Dear all, I am using confocal leaser microscope for calcium imaging. Last couple of days I have faced difficulty to get the cell responses, even though I did not get the response of ATP. I am using HEK293T cells and transfect plasmid DNA using ScreenFect (Wako). I would like to see the cell responses of different agonists and mixtures of known agonists and putative antagonist. I have always used 5 µM ATP for normalizing cell activity. But, last few days I did not get any response from ATP. Can you please suggest me about this problem.
Thank you in advance.  
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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
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I have the std curves alizarin red solution using CTC
I also have the total protein cantent values to which Ca conc. has to be normalized.
eg: The concentration of ARS in mg/ml = 3.8766
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thank you..:)
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I am looking for a way to study the function of synapses on neurons in culture without having to do e.phys.
Calcium imaging, e.g. FLIPR?
Thanks!
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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!
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I am using MCEC (Mouse cardiac Endothelial cell line) and using Methylglyoxal as agonist. I would like to know what could be possible ways to explain this phenomenon. I have also tested and it is not an artifact as agonist is not interacting with FURA dye. 
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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
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Hello, I'm studying memory in crockoaches and I would like to see the calcium imaging influx. Does anyone know a protocol using Fluo 3/4 in vivo/in situ? I can't do cell culture. I didn't found a good paper talking about it. Thank you for your attention. 
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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.
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Hi again:
I am trying to optimize a protocol for imaging calcium dynamics in a cell population in the dentate gyrus and am facing some trouble. I made a similar question and got 1 answer but I guess a new one attracts more people and I am going to be more specific.
I am making coronal acute slices and staining them with a calcium indicator, but when I mount my slice I don't manage to focus it under the 40x objective or sometimes even the 20x one. I place my slice onto a glass coverslip that fits into a chamber where ACSF flows through.
I think the problem is the dentate gyrus detaches from the coverslip, maybe the orientation makes it easy. I have tried attaching the slice with PEI and poly-L-lysine (PLL, coating the coverglass over night) and putting a small piece of metal on the slice but it is not working for me. I have read that PLL doesn't work so well with tissue and gelatin sticks better, but I am also afraid that my slice can wrinkle and get stuck too soon. Any ideas?
Thanks in advance and kind regards,
Roberto
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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
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I have been loading Fura2 at room temperature, 30minutes after washing steps I measure with FACS Calibur. I do not see any change comparing with unloaded cells. I will be grateful to have a working protocol.
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5 uM loading for 30 minutes at 37 C. Wash twice.
What lasers/detectors are you using? Have you used a positive control?
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I am woking with calcium sensor protein and I am facing trouble with aggregation. I want to use PEG as a stabilizing agent, but I don't know if I can use PEG in Calcium binding studies or not.
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Can you try BSA instead?
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Hi, I'm new to calcium imaging but have been doing electrophysiology (whole-cell patch clamping for a few years). As our imaging system does not have a perfusion system and no carbogen gas available, we are not able to perfuse the brain slice with oxygenated ACSF during imaging. This of course will impact on cell survival due to changes in pH and low oxygenation. Otherwise brain slices were maintained in oxygenated ACSF prior to imaging.
Does anyone have a protocol for doing calcium imaging without the slice being oxygenated? i.e. a recipe for a buffered solution that do not require oxygenation during imaging?
Thanks!
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The carbogen bubbling is essential for for pH buffering via the sodium bicarbonate of the ACSF. Organotypic brain slice cultures and primary cultures live happily for weeks at  atmospheric oxygen levels when buffered by HEPES, and the oxygen component of the carbogen may not be as important as the CO2. 
HEPES is frequently used to buffer ACSF in absence of carbogen, and I think it should readily work for you without perfusion, at least for some tens of minutes and probably even longer. Pay attention that you dont have too much evaporation of ASCF during the recording, which may increase the osmolarity of the solution (basically dont have too low a volume in the imaging chamber).
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I am isolating mitochondria from cell lines and then I do an assay to check for mPTP opening. I use a dye sensitive to calcium and check for uptake and release by mitochondria. However, sometimes the OD reading goes down fast and then increases and then goes down slowly again. Has anyone encountered that before? Any suggestions on how to fix it? Thank you.
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 Full text of above article
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According to the companies Description, BAPTA is Ca2+ chelator exhibiting a 105-fold greater affinity for Ca2+ (Kd = 110 nM) than for Mg2+. After BAPTA complexes with Ca2+, the UV spectrum shifts from 254 to 279 nm at pH >6.0. Useful for spectrophotometric monitoring of extracellular Ca2+ levels, especially transient phenomena, because of its insensitivity to pH.
I have used 50mM Tris Hcl (pH:9) for preparing 10mM BAPTA solution. then I used this buffer for determining a standard curve at different ca2+ concentration (10^-8 M to 10^-4 M). but I only saw a sharp absorbance at ~230nm which didn't change significantly following Ca2+ addition. Is there any body who worked previously with this compound? any advice will be greatly appreciated
(please note that I am trying to concentrate free Ca2+ ion by an indicator to determine ligand+protein binding)
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The standard solutions would not help anyway since I guess your normal working extracellular medium Ca would be in saturating range for BAPTA. The lack of spectral effects between 10 nM is 10 mM is and enigma indeed. Then the question is, what was the source of your Ca standards. If you made them yourself, in well could be contaminated up to micromolar range.
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Hi again:
I am trying to optimize a protocol for imaging calcium dynamics in a cell population in the dentate gyrus and am facing some trouble, so I resort to this community again.
I am following pretty much the protocol I link, but the attaching process with PEI is not working for me. I need to attach acute slices after staining with the calcium indicator to a coverslip that fits in a perfusion chamber suitable for confocal imaging. I read that coating coverslips with poly-l-lysine is the usual solution, but I also heard slices need some time to attach and I need to perform the imaging on the same day I get the slices (mostly because of the age of the animals). Could you help me with your experience?
Thanks in advance!
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Hey Roberto,
I could not access the link you provided for the protocol. However, I coat my coverslips  with Poly-L-lysine (0.01% solution, Sigma) and let it dry under laminar flow. Once dried, I put mouse brain slice (sagittal or coronal, up to 400-micrometre thickness) on coated coverslips. It sticks very well and almost instantaneously to these coated coverslips.  I use them for electrophysiological recordings but you can give it a try. 
Hope it helps !
best luck,
Nand
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Hello everyone. I am trying to optimize a calcium imaging protocol in a population of the dentate gyrus and using Fluo-4 in the first steps as it was the indicator available. For the moment I have observed some stained cells in an acute slice with a fluorescence microscope and I am moving forward to the confocal in order to see changes and of course have more resolution. My way to visualize the population is transgenic GFP expression, so until the moment I optimize the protocol with another indicator I would like to know if someone has experience distinguishing between GFP and Fluo-4 spectra in a confocal setup.
Thanks in advance!
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Difficult problem. See
Wilson JM, Dombeck DA, Díaz-Ríos M, Harris-Warrick RM, Brownstone RM. Two-photon calcium imaging of network activity in XFP-expressing neurons in the mouse.  J Neurophysiol. 2007 Apr;97(4):3118-25. Epub 2007 Feb 15.
PMID: 17303810
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I would like to raise the free intracellular Ca2+ concentration to within the range of 2-5 uM. Many researchers use Ionomycin or Thapsigargin to raise intracellular Ca2+, however, based on the published Ca2+ imaging data it looks as though the free Ca2+ rises to a few hundred nM at most.
Is there a way to augment the effects of either Ionomycin or Thapsigargin? Since Ionomycin is an ionophore, perhaps increasing the extracellular Ca2+ concentration during the application of ionomycin would help? Are there other reagents (besides ionomycin and thapsigargin) that would better suite my need?
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Hi Gary,
HEK cells have limited surface calcium channels and you don't have many choices to raise intracellular Ca2+ levels other than using a Ca2+ ionophore like Ionomycin or blocking SERCA with Thapsigargin as already suggested by the colleagues. If you are afraid that the intracellular Ca2+ levels are not high enough using this approach (I believe not high enough compared to controlled cells), you may want to diminish Ca2+ levels in your control cells using BAPTA-AM for example, so that the the difference in Ca2+ levels between your "control" condition and 'high Ca2+" condition is increased which should allow you a better comparison. Not perfect situation but an alternative.
Hope this helps. Norbert 
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