Questions related to Calcium Imaging
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!
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?
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.
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?
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
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
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!
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.
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 ?
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.
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?
Which approach is better for in vivo calcium imaging in the brain with miniscope: image in virus-injected (AAV GCAMP6) or transgenic GCAMP6f animals?
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!
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.
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.
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.
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
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.
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 ?
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?
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?
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?
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.
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!
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).
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 !
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
Please provide solution that is readily available. Also what is the downsides of each of them when comparing to electrophysiology?
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:
- 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.
- 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!
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.
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.
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!
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.
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
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.
PhD student at IGBMC,
Translational Medicine and
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?
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?
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
I'd like to have some advices to perform calcium imaging in embryonic neuronal cultures.
Thanks to anybody who will help me
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?
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 !
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!
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
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 ?
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?
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.
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.
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,
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
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?
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 !
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.
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!
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.
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.
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.
- 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!
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?
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.
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
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?
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.
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.
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,
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.
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.
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?
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.
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)
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!
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!
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?