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According to the World Health Organisation, ‘long COVID’ can be defined in the following way: “Post Covid-19 condition occurs in individuals with a history of probable or confirmed SARS CoV-2 infection, usually 3 months from the onset of Covid-19 with symptoms and that last for at least 2 months and cannot be explained by an alternative diagnosis.”
It is estimated that 1.2 million people in the UK were reporting long Covid symptoms in the four weeks up to 2nd October 2021 by the Office of National Statistics (ONS), about 1.9% of the population
Imperial College London have reported data about the symptoms of long COVID in their COVID Symptom Study, identifying two main groups of symptoms.
Some reports have described symptoms with similarities to Myalgic encephalomyelitis (ME), also known as, chronic fatigue syndrome or ME/CFS.
There are many multifactorial and complicated mechanisms involved in the pathogenesis of COVID 19 and other neuropathology symptoms have been described, such as, pain, dizziness, headache, dysgeusia, or anosmia and flacid paraparesis to more serious symptoms including stroke, Guillain-Barré syndrome (GBS), acute haemorrhagic necrotising encephalopathy, meningoencephalitis, and cerebral venous thrombosis.
Many patients experiencing long COVID symptoms are ‘flying under the radar’, without much medical intervention, but may have significant deficiencies affecting their ability to perform at their usual state of health.
If mild to severe versions of Guillain Barre were a factor in the long COVID symptoms, should they undergo investigations for Guillain Barre and this be a new approach in their treatments? Are long COVID patients being assessed with GBS or other neuropathology variants in mind?
Further, what can we learn from the MERS outbreak about neuropathy and cerebrovascular disease?
Should we be performing nerve conductivity tests on long COVID patients and investigating autoimmune mechanisms to improve appropriate treatment strategies?
Journal of Molecular Neuroscience (2021) 71:2192–2209 https://doi.org/10.1007/s12031-020-01767-6
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The following RG link is also very useful:
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1. How Long does it Take for Tamoxifen to activate Cre and knock out the gene of interest
2. What would the dosing regimen be for rats/mice?
I am designing an experiment where I will be knocking out 5-HT2A receptors in a cell type-specific manner before psilocybin administration to determine the necessity of 5-HT2A receptors in psilocybin's neuroplastic effects.
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Which cells do you want to reach ?
It is extremely important to know the target location. e.g. blood-brain barrier.. 1-2 dosages are sufficient, intracerebral (e.g. astrocytes, neuron) can take a while.
Depending on your animal facility you will have difficulties due to quarantine time
of the tamoxifen.
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I am curious about two specific things:
- Why do pseudounipolar neurons have one axon (as opposed to a dendrite + axon like multipolar neurons)? How does this structure reflect sensory function?
- How do potentials propagate through the axon? Since there is no axon hillock for summation, does that mean no summation occurs? Is there still a threshold potential that needs to be met? Or does every graded potential get transmitted through the axon?
Can someone familiar with any of these questions help out or provide a resource I can refer to?
Thank you!
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Hi Sehej
Pseudounipolar pattern of sensory neurons acts as low-pass filtering, potentially regulating sensory information reaching the spinal cord. Thus, by impedance mismatch between membrane point in the vicinity of T-junction, this has been recognized as a site where spike propagation may fail.
As for action potential propagation through the axon, this is a more broad question. Actually, in a really simplistic summary, sensory neurons have "transductors" (specialized proteins that converts physical energy - thermal, mechanic, chemical - into electrical signal) in their peripheral end; these transductors creates a "generator potential", according to specifical thresholds. If, and only if, these generator potentials reach some specific area in the membrane with a larger density in voltage gated channels (as Na channels) within these second step threshold, an action potential (spike) is generated and conveyed until the "T-junction" filter described above.
Some references for better comprehension
1 -Al-Basha, Dhekra, and Steven A. Prescott. "Intermittent Failure of Spike Propagation in Primary Afferent Neurons during Tactile Stimulation." Journal of Neuroscience 39.50 (2019): 9927-9939.
2 - Sundt, Danielle, Nikita Gamper, and David B. Jaffe. "Spike propagation through the dorsal root ganglia in an unmyelinated sensory neuron: a modeling study." Journal of neurophysiology 114.6 (2015): 3140-3153.
Regards,
Tiago Avelar
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I was wondering if I fear condition my rats, and then train them in Morris Water Maze (spaced training), will the rats show weaker or no long term spatial memory? I tried to look for relevant literature, but could not find many papers directly addressing this sort of question. If I missed them, could someone send me those papers? Thank You.
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From your question it's not really clear, do you aim to test such a hypothesis or to avoid unwanted effects of one task on another?
Fear conditioning in rats can be quite stressful, so the effects of stress modulators (corticosterone, noradrenaline) can definitely affect learning in the next task. But the effects will depend on shock intensity, the timing of each task etc.
If you could be more specific I'd love to provide more literature recommendations.
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It is my first time to measure the level of ROS. especially in brain both in vivo and in vitro. I am not quite sure that there is an recent method which is high accuracy to investigate the ROS level in brain. So, is there anyone have an experience on this kind of experiment? Could you please give me suggestions?  I will very appreciate for your kind. Thank you.
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ROS has a half-life of milliseconds and thus analysis of fixed or frozen material may not reflect the in vivo situation.
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Hi! 
Please let me know if you or your lab in Europe have  Ndnf-IRES2-dgCre-D transgenic mice. I will be extremely grateful!
Many thanks!
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We do have this line and currently characterizing it.  This is the line from Allen brain institute.
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Can we inject combination of AAV virus to brain in same time? I am planning to inject AAV2 as well as AAV9 as a co-injection in mice.... any thoughts?
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Hi Dilshan, you probably have thought about this already, but just to remind you that since each serotype has its own host tropism, they might not co-transduce the same tissue or cell type with the same efficiency. If you are stereotaxically injecting the virus then probably this doesn't matter, as AAV2 and AAV9 are both good serotypes to use in CNS. Also, if they transduce the same cell, sequences with homology might undergo homologous recombination at a low but detectable rate. 
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Dear colleague,
I would like to stimulate my recorded cell (whole-cell, pyramidal cells in hippocampus) with a pulse train of light during 600 ms. In the middle of those pulses of light, I would like to stimulate electrically the axons (0.1 ms, 1 pulse). How can we "superimposed " the two stimulations with clampex 10 ? For now I found how to build the protocole with one stim after the other. But I would like to stimulate electrically while the pulse of light is ON.
My electric stimulator is branched on the Out#0 and the LED is branched on the Out#1 of a Digidata 1500A.
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Under the waveform tab you set up three waveform epochs, the first one is just light, the second one is light plus stimulus and the third one is just light again.
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Dear all,
I am recording sEPSC in layer V neurons in mouse mPFC, but I found some abnormal responses showed as attached pics. They look like epileptic discharge in presynaptic neurons,  is there any e-phys expertise can tell me what was wrong with my recording?
I held the neurons at -70mV, and add picrotoxin 100 uM to the ACSF.
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Hi, what do these look like under CC? I suspect they are the compound EPSCs that cause paroxysmal de polarization shifts. I think this is a result of ictal network activity you're getting from applying picrotoxin to the entire slice.
If you're using it to disinhibit everything on purpose, you are in a tricky place... 
If on the other hand you just use the picrotoxin to block sIPSCs, I'd recommend local application (ytube) to avoid the paroxysmal activity that's likely causing these giant currents! 
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Dear colleages,
I'm trying to evoke LTP in CA1 of 2-3 months old mice using a TBS protocol. Up to now, I've tried the two following ones:
a.- Bursts of 4 pulses at 100 Hz, given 10 times and interspaced 200 ms. 
b.- The very same stimulation protocol but applied three times with 20s interval.
Up to now, I have just tried in controls -finding LTP in almost all of the slices-, but I am about to start doing experiments in transgenic mice and I would like to know which protocol may be better in my condition. Which are the benefits and caveats of each of these protocols? I guess the three-times stimulation protocol is stronger, but I find in most papers the authors use the one-time stimulation.
May someone contribute with some hints?
Thanks in advance;
Sergio
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Dear Sergio
it depends a little on what exactly you want to investigate. Both protocols are pretty strong. They will elicit (close to) the maximum possible potentiation. The second protocol will probably be strong enough to cause L-LTP, the very long lasting (3 hours), protein synthesis dependent form of LTP. So, if you are interested in those processes, that would be the protocol to go for. However, if you have a mutant that leads you to believe the difference might not be in the protein synthesis dependent phase or in the maximum amount of potentiation, I would go for another protocol.
I would suggest to use two protocols. One using only two trains of 4 pulses at 100Hz spaced 200ms. This is the absolute minimum. It will be on the threshold of evoking potentiation. Any very subtle difference between mutant and wild-type would probably be picked with this protocol. The other I would use the more standard 4 trains of 4 pulses at 100Hz spaced 200ms. Record for 1 hour after induction, unless you are interested in the L-LTP as mentioned above.
good luck and do not hesitate to ask follow up questions
Nils
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Two papers are using the method.
One attached and the other cited here: "Single rodent mesohabenular axons release glutamate and GABA" Root et al  2014
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Supplemental figure 3 and supplemental text from "Cho JH, Deisseroth K, Bolshakov VY. Synaptic encoding of fear extinction in mPFC-amygdala circuits. Neuron. 2013 Dec 18;80(6):1491-507." have a very nice explanation of the logic behind using TTX and 4-AP isolating monosynaptic inputs. They also show a positive control in which disynaptic inputs are not rescued by 4-AP. Here is a link to their supplemental materials: 
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I'm having issues with tissue processing of mouse brains and spines. Both tissue types appear white and opaque (milky) - especially once mounted and dried on slides.
Brain tissue also appears to expand and disintegrate after a few seconds on floatation (at room temperature and at 42 C). The wax stays intact.
My protocol for these formalin-fixed tissues (~3mm thickness) is:
1 hour - 50% alcohol
1 hour - 70% alcohol
3 hour - 95% alcohol
1 hour - 100% alcohol
1 hour - 100% alcohol
1 hour - 100% alcohol
2 hour - 100% alcohol
1 hour - xylene
1.5 hour - xylene
1.5 hour - xylene
2 hour - Paraplast Plus paraffin (no vacuum)
2 hour - Paraplast Plus paraffin (no vacuum)
I'm using reagent alcohol (90% ethanol, 5% methanol, 5% isopropanol).
They will be stained for H&E and luxol fast blue.
Is this due to either insufficient dehydration or clearing? Does using reagent alcohol instead of ethanol make a difference?
Any advice would be greatly appreciated!
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I think the protocol looks ok. Also the use of reagent alcohol (ethanol+methanol+isopropanol) should be ok, as fas as it is water-free in the 100% alcohol. 8 hours in sum in high-percent alcohol seems rather long for small brain-samples, but should not result in insufficient dehydration.
Is the surface of the paraffinblock "wet", slushy and does it sink in the center until the next day? These would be signs of residual xylen or ethanol in the tissue. Then one can put the tissue again in molten paraffin for an additional day and then cut again.
Check, if the reagens in the processor are fresh and in the right order.
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What would be the specifications of BCN cables (for connections between Axon Digidata 1440A and Multiclamp 700B apparatus) for carrying out appropriate in vitro extracellular recordings?
I'm afraid the coaxial BNC cables I'm using are not enough to avoid noises.
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Hi Alexandre,
I agree with Zuner that the internal resistance of the cables is so low as to be not important. I suspect, however, that there are quality differences in the cables themselves and cheap ones should be avoided. I presume you are sure that the noise is due to these cables and not some other source? If all else fails you might try wrapping the cables and the outside (earth) connectors in a double layer of aluminium foil, it might help to have some extra shielding.
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I want to contruct a primer for assay of scavanger receptor marker MARCO , or CD163 for rat animal model. So my question is that how to consttruct a primer for this ?
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so are we talking PCR based detection? If that is the case check out taqman probe availability for this receptor. I am pretty sure they provide kits where you can design whatcha need.
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Does anyone works with this molecule in ratt tissue? I am trying to find the best commercial antibody in order to detect CB1 receptor by immunofluoresce and western blot. 
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Hello,
I would recommend a free antibody database available at labome.com. For anti-rat CB1 antibodies for IF or WB you can check the following link: https://www.labome.com/review/gene/human/CNR1-antibody.html. Invitrogen has rabbit polyclonal CB1 antibody (ABR, PA1-743) which can be used in immunohistochemistry and WB  on rat samples (ref: López-Gallardo et al. J Neuroendocrinol. 2015). Also, Abcam rabbit polyclonal CNR1 antibody (Abcam, ab23703) was used in immunohistochemistry on rat samples at 1:300 (Meng et al, Int J Clin Exp Pathol. 2014). You can see the description of these antibodies and references following the provided links.
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Hello,
I am trying to optimize our Olig2 immunohistochemistry staining protocol for PFA fixed mouse brain. I found a very helpful website that reviewed several recent publications and found people used concentrations from 1:100 to 1:500 on mouse brain, and I am planning to try several of them (https://www.labome.com/review/gene/mouse/Olig2-antibody.html) . However, on the abcam website they advise to perform heat mediated antigen retrieval with citrate buffer pH 6 for this antibody. I was wondering if anyone tried antigen retrieval for Olig2 IHC?
Otherwise, if you have experience with antigen retrieval using a microwave, could you please advice the steps? I found a protocol suggesting placing tissue sections on slides, in citrate buffer (pH 6), and boiling for 20 minutes, whereas another protocol suggested boiling for 7 minutes.
I was also wondering, after antigen retrieval, what is the best way to continue the staining protocol having the sections already on slides: i.e. the incubation with the primary antibody over night etc? I have only ever stained with the tissue inside well plates, not on slides.
Thank you,
Carola
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For microwave antigen retrieval, I use a glass Coplin jar. Put your slides in, fill with citrate buffer, no lid. Microwave on 100% power until just starts to boil; ~1 min or so. Then reduce power to 30% for 10 min. Keep an eye on it. You don't want a rapid boil. If you need, add more citrate buffer. After boil, take them out and cool on bench for about 30 min. Wash with buffer a few times then go forward as normal. I use a PAP pen to draw a circle around my tissue and then block and stain within that circle. Block 1 hr with PBS + 3% BSA and maybe a little rabbit or goat serum if you have it, then O/N 4o with primary. Next day wash and then secondary Abs for 1 hr RT. Wash and go forward with whatever method of revealing you are using, i.e. fluorescent or DAB, etc. Hope this helps.
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I have around 8-10 mg tissue. I don't know whether it would be suitable to use TRIzol method.  If anybody has isolated RNA from this much quantity of tissue using TRI, I just want to know if their is any additional process to follow to ensure that we dont lose the tissue?
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Hi Pooja,
Your tissue (not RNA) is lost when you throw it in the TRIzol. 
I routinely isolate tissue from 5 day old zebrafish which have a weight of <2 mg. In these fish there is a lot of RNA present. However when you are dealing with e.g. muscle tissue, 8-10mg tissue is very difficult to get normal RNA out. Fresh brain tissue should be fine.
So it is all about what kind of tissue you are dealing with!
One of the best ways to isolate this amount of RNA is through TRIzol treatment. However you should use a carrier like glycogen or glycobleu (max 1/10 of  your final solution where you store your RNA in).  You add glycogen before pelleting the RNA. If you want i can share you the protocol I use for the fish.
Good luck,
Bart
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I have transduced different cell lines with a Cre and Cre Reporter (GFP/RFP colour switch) lentivirus (Gene Target LVP339-PBS for Cre and Gene Target LVP460-Puro-PBS for Cre Rep). When I co-culture those cell lines ‘immediately’ after the transduction, I see a good amount of recombined cells (around 80 cells in a well of a 96 well plate when 20k cells were seeded). However, the number of recombined cells decreases after each passage, resulting in only single recombined cells after 4-6 passages, depending on the cell line. The reporter line does work when I add TAT Cre to the cultures, so this is not the problem. I have cultured the Cre cells under puromycine since transduction and I have sorted the reporter cells for GFP expression using FACS.
Has anyone experienced the same or knows how to get around this?
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We are using a CMV promoter and differnet human and murine cancer cell lines. I think it is unlikely that the promoter is downregulated.
We have two different viruses, one with a puromycine selection marker and one that co-expresses GFP. We have tried both, antibiotic selection and FACS sort for GFP positive cells. However, we always end up with a decrease in Cre expression after every passage...
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I am trying to compare yield of fresh vs FFPE and the measure relative quantity by RT-qPCR of housekeeping gene. I can upload the protocol I am using right now if anyone is interested in sharing their opinion too, but what I noticed is it doesn't involve any sort of homogenization like the most fresh tissue RNA extraction protocols do...  I have attached a link of the protocol I am using below.
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Try to use Qiagen RNeasy Kit for Purifying total RNA from FFPE which i used before and gave me a good results .
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Hello! Has somebody used the ED1 clone of the CD68 marker of microglia in rat brain homogenates? I'm planing to use it as a marker of "activated" microglia in WB
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We use it frequently for IHC on mouse and rat frozen brain sections. It works very well to mark activated microglia/macrophages. I have not tried it for western.
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I would like to measure dopamine release in anaesthetised rats with continuous amperometry, what anaesthetic is most suitable for this experiment considering that I cannot use urethane? Thank you!
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There is a publication from Mark Wightman group comparing DA overflow (release and reuptake) in the caudate after stimulation of the DA neurons during anesthesia and in the same awake animals. They used FCV. As far as I remember release was not much affected, but uptake was a bit slower after chloral hydrate. To make amperometry less noisy you may use much more sensitive carbon fiber electrodes as for example electrodes manufactured from 30-32 micron fiber. Indeed, this will not improve selectivity.
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I want to measure brain damage in EAE mice but i dont hve any experience about that
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I can send you LFB protocols on paraffin or frosen sections, as you prefere. It worked in rat and cat brain. But I am really curious how you will quantify the brain damage. And why don't you try Myelin Basic Protein Staining? LFB stains not only myelin. MBP is more specific
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I am trying to image my zebrafish embryos from 0-120hpf and am struggling with suitable mounting without damaging the specimen. Does anyone have a good protocol or own experience for it? 
I am using Leica TSC SP8 motorised inverted DMI 6000 microscope.
Thank you
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Since you are using an inverted microscope, make sure there is a cover slip at the bottom - do not use the usual microscope slides. You can make a hole in a small plastic petri dish and attach a cover slip to its bottom surface. Multi-well plates or chamber slides work fine too, since they are designed to work with inverted microscopes. I had good results with low-melting point agarose, as suggested above. It is easy to change the surrounding medium with it without displacing the embryos (particularly useful if you are planning on using a clearing solution). However, if your embryos are alive, the resistance of the agarose can prevent normal development. Some people use methylcellulose, but one needs to be careful to prevent any evaporation of water from it. It dries out quickly.
Otherwise, here is a protocol:
and there is also this, if it helps:
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If I make a sciunit TestSuite that only consists of two types of tests (see below), the NEURON simulator is called at least six times, where as I would have only thought two invocations are necessary to obtain error values. Are the additional invocations of NEURON because neuronunit is generating predictions in order to produce a data-derived (see hyperlink) model as opposed to a simulator derived model? I think that NeuronUnit first uses a simulator in order to produce a data derived model and then it creates an error from this data derived model.
Is a data derived model really different to an empirical observations (other data that may originate from Allen Brain, or neuroelectro for example)? If the data derived model is directly related to the empirical observations, I am left confused about why neuronunit makes so many additional calls to the NEURON simulator.
tests += [nu_tests.RheobaseTest(observation=observation)]
test_class_params = [(nu_tests.InputResistanceTest,None)]
The file below gives the greater context of the above code statements.
Also note that neuronunit is only called with test which is defined as
test=tests[0]
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I suspect the Rheobase test, when searching for the rheobase current, will try several injection current values until one elicits a spike. To ensure there is no contamination from previous current injection, it restarts the sim from scratch.
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A thought experiment I would love feedback on:
If the membrane conductance of a neuron was increased (ions could more readily pass back and forth), but without complete breakdown of the resting membrane potential (i.e. still able to undergo spontaneous firing). What would the wave of voltage verse time as measured from a stationary electrode look like compared to a typical neuron?
My assumption is that the resulting voltage 'spike' formed by a depolarization event would be shorter and wider than that of a typical neuron, because the decreased magnitude of baseline potential would result in an lower change in the voltage (smaller amplitude) and the increased conductance would result in a longer interval to restore resting potential (wider spike).
Is my thinking correct on this?
Thanks.
I
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dv/dt = I/C seems like a good equation to start with.
Secondly, how is what you describe different from a subthreshold EPSP or IPSP?
Thirdly, you didn't comment on what the reversal potential for this conductance is? If it is the same as the resting membrane potential, then there will be no voltage "wave".
Finally, and most directly, assuming the reversal potential of the conductance is not equal to rest, if the conductance ends begins and ends instantly, the the charging and decharging of the membrane will follow an exponential with a time constant equal to the membrane resistance multiplied by the membrane capacitance. (The beginning and termination of the conductance do not have to be instantaneous, just much much faster than the time constant of the membrane).
Or are you talking about what an increased membrane conductance does to an action potential waveform? The answer is "not much", as the conductances involved in generating an action potential are orders of magnitudes greater than typical resting conductances, assuming the conductance you are operating isn't far from rest. I've attached a simulation of a spiking neuron with a resting conductance of about 5 nS, driven to spike via a 2ms long somatic current step. I then introduce a 2 nS conductance with the same reversal potential as the resting membrane potential and increase the size of the current step to produce spiking at almost the same time. As you can see, the shape of the AP is not noticeably altered.
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Hello everybody,
I want to use a Cre-Lox mouse model to study the behaviour of glial cells during a Parkinson-like state. In our in vitro model, we take glial cells from newborn mice to perform our primary cultures.
I am wondering whether I can treat first these Cre-lox mouse and afterwards put newborn glial cells in culture ? Or is it better to take glial cells from Cre-Lox newborns before treating them "in vitro" in the plate with Tamoxifen ?
Thanks in advance,
Have a nice day,
Best,
Tony Heurtaux
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As far as i am concern the breeding of two parent one carrying gene to be knock out flanked by loxp and other parent carrying cre gene under the control of tissue specific promoter. The offspring therefore carry both the gene. So the gene will be knock out only the tissue that is purely determine by cre gene promoter. if your cre gene is exclusively express in in glial cell it will knock out loxP gene gene in glial cells. Therefore treatment condition is purely base on what factor determined the activate cre protein through cre gene promoter.
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Is it possible to analyze the neuron morphology (marked with anti-MAP2) by optical density in image J? I want to measure the cellular projections of neurons which were exposed to toxicity.
Thank you.
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You can also use Neurolucida 360 for 3D volumes (tissue) or NeuronJ (Free ImageJ plugin available here: https://imagescience.org/meijering/software/neuronj/ ) for 2D images (culture)
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I am running western blots on adult rat brain (white matter) and at the moment I am getting too many bands and it would be good to use negative control to see whether of those bands are full length of NRG1 and c-terminal fragments. I could use NRG1 knockouts but currently we don't have any, so I am looking for alternative. Also, it would be good if you could advice me on a good primary antibody. The one I am using is from Santa Cruz (Neuregulin 1α/β1/2 (C-20)).
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No but you can check if the immunogen sequence of Human NRG1 is retrieved in mouse. 
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Hello,
I am planning a ChIP-Seq experiment from human brain samples (for example, 10 case and 10 controls samples). I was wondering, since I am new to ChIP-Seq, will I need to add the input samples for each sample into my library as a control? So when performing ChIP-Seq, do you need to sequence both the ChIP-DNA of interest and the input from that same person for every sample you are interested in?
Any help on this matter will be greatly appreciated. Thanks.
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I agree with Ian. We have frequently run ChIP-seq experiments where we pool all Inputs and only sequence the pool instead of each independent input. Optimal of course is sequencing each individual but most times the pool is sufficient. I would also recommend to always run a control qPCR on a small aliquot of your IP with positive and negative control primers for your target of interest before creating libraries.
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Many kits are declared just for plasma or cell culture samples. Thank you!
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Hi Katerina, I see that others are already recommending the Biosensis ELISA kit. A recent paper from Enrico Tongiorgi's group published in Nature Scientific Reports compared most kits and found the Biosensis kit to be superior in several respects. Details are on the Biosensis data sheets and protocols. However, there are no kits that have been fully validated for CSF. Biosensis scientists have begun to examine CSF so you might want to contact them directly. Please note that I have been involved in the development of this Biosensis kit as an advisor, and that I am also declaring that I a shareholder in the company. 
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How to prepare a single cell suspension from without disrupting the membrane integrity for ADP/ATP ratio measurement?
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I want to prepare the single cell suspension from brain especially hippocampus to measure ADP/ATP Ratio.
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Looking for any normal brain cell line.
Can collect if in west mids or pay for postage if further.
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I'm afraid there is no such thing as a "normal brain cell line". I think you'll have to be a little more specific about what you're looking for....
best,
Agnete
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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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Dear all, I have run my tPA zymography protocol for more than 1 year without problems. Last week it stopped working. I made all new buffers and still is not working. Anyone had the same experience?
Here is the protocol: 12.6% SDS gel with casein and plasminogen (both substrates are stored in aliquotes at -20 or -80C). Running at 110V in the cold room. 2x30 min rinses with 2.5% triton-x solution, 3x10 min rinses with water. Incubation in glycine/edta buffer pH 8.3, coomassie blue R250 staining overnight, destaining, rinses in water.
Thank you in advance!
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Dear
Please check the following publication to find another interesting informations: Use of Gel Zymography to Examine Matrix Metalloproteinase (Gelatinase) Expression in Brain Tissue or in Primary Glial Cultures. Harald Frankowski et al. 2012. Methods Mol Biol. 2012; 814: 221–233. Directlink: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3670093/
Hoping to be helpful
Marius
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I am currently perfoming multiple brain microinjections in male SD rats over a number days. Unfortunately, I am finding myself needing to replace cannulae obturators (currently using 33G wire) almost daily as the rats are very skilled at removing them. As a result, many cannulae are becoming blocked. Can anyone recommend ways to construct obturators that may reduce the amount being replaced and ultimately, the number of blocked cannulae? Thank you in advance. 
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PlasticsOne (http://www.plastics1.com) make a nice set of guide cannulae, with stylets and screw-on caps. Blocking is still occasionally a problem, but we found that if you take the cap off and gently move the stylet in the guide cannula every day, the risk of blocking is greatly reduced.
Best wishes
Chris
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Dear all, 
We do a lot of cFOS IHC stainings in our lab and so fare we have been using a great antibody "rabbit-anti-cFOS" (at a dilution of 1:10'000). Unfortunately this antibody is not available any more. 
Since a coupple of month we are struggeling to find a new antibody that stains mouse brain tissue such as the AP (area postrema), NTS (nucleus tractus solitarii) or ARC (nucleus arcuatus). 
We either get very weak stainings or no staining at all with the antibodies we're currently testing. Also we have to use them at much higher concentration (such as 1:100/200) as compared to the antibody before (1:10'000). 
Thank you for your answers, 
Fabienne
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 Dear Fabienne,
I'm doing lots of Fos staining in mouse brain, mainly in the hypothalamus (including the ARC).
I always used the two following ABs with high staining quality :
1/ DAB staining: rabbit polyclonal anti-c-fos antibody Santa Cruz – c-Fos (4): sc-52R, dilution 1:2000
2/ Fluorescence and DAB: rabbit polyclonal anti-c-fos antibody anti c-fos (Ab-5) (4-17) pAB rabbit, dilution DAB 1:15000 2xON, dilution fluo 1:5000 2xON
Hope it will help and let me know if you need further details,
Bests,
Olivier.
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Has anyone had success in detecting NGF, BDNF, NT-3/4, IL-6/8, or IL-1B in neuron culture media by Western blot, or are ELISAs required? If Western blot worked for you, what antibodies and concentrations did you use? Thanks!
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Hi Katherine,
To my experience measuring BDNF, IGF-I and GDNF from cultured neural progenitor cells, I would prefer using ELISA kit with good sensitivity and specificity. WB could detect them also but it requires higher amount of those proteins in your culture media since its sensitivity is lower than ELISA. 
Narisorn
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We have been performing a thioflavin-T dose-response curve on alpha-synuclein fibrils with very consistent results for some time now.
Recently, we received a new batch of fibrils produced by the same method as before, but by a different person as the original person has moved elsewhere. Since then we have not been able to get the same dose-response curve.
If you look at the attached graph, curve E7 was obtained using the original fibres and curves E16 1 and E16 2 using the new batch. Does anyone have any idea what might be happening?
I have had a lot of help with this experiment on here, so I hope someone can help with this question too.
Previous discussions on this experiment are below.
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I forgot to mention in the question that the concentration of the fibrils is kept constant and varying concentration of thioflavin-T is used to obtain the curve.
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Recently, I'm doing the rat and mouse cortical neuron culture, using PEI coatin g 48-well plate. I seed 8-100000/well,but it's easy to clustering or dead in 3 DIV. The growth of neurons were not equivalent between different wells. And mouse cortical neuron seems more fragile than rat. Any difference between them? How can I manage to culture cortical neurons for toxicity test? 
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For toxicity you always want your agent to be the cause of death, not the plating technique.  If your well-fed neurons are not surviving for at least two weeks without toxin, then your culture system can be improved.  As Maria (hejsan) explained, density and coating are key...
Density - If neuronal cultures are plated too densely, they will cluster, and if they are not dense enough they don't survive very well either.  A general safe-range formula I use (for mouse cortical) is ~ 40,000 to 60,000 cells per cm2 of your coated surface (in the case of an 48-well plate with 6mm coverslips = 0.3cm x 0.3cm x 3.14 x 60,000 = 17,000 per COVERSLIP on the high end.  So, it may be that your density is a little too high.  It sounds like you might be plating directly into a coated plastic plate though, so the 48-well dimensions are a bit bigger and cell number should more like 50,000 per well and closer to your current range.  Try once just adding fewer cells, and see where that takes you.  If that does not do the trick, then continue reading...
You should consider using glass coverslips.  The inconsistency between wells might be due to the cell suspension not being transferred evenly, which would be more an issue with smaller volumes of 48-well plates.  So try glass coverslips that have a bigger surface (12mm in a 24-well plate) and you won't go blind - Maria's suggested cell density is a good range for this size.  
Coating - For cortical neurons, most coat round glass coverslips with Poly-D-Lys (I use 50ug/mL), followed by Laminin (10-20ug/mL).  I am not sure how PEI coating works, and don't want to be bias to the old school methods when I haven't tried the other.  In case you want to try what old people use, PDL/Laminin coating can give consistently adherent cultures.  
>Add minimal (50uL or less) Poly-D-Lys to each coverslip as a surface tension dome resting on the glass, instead of flooding the entire well around the coverglass just to get the meniscus above the center areas. Incubate this for 1-2hrs at 37'C
> Then pipette in Laminin directly into your PDL dome to the appropriate final concentration (this will also save you on Laminin).  Incubate for 1 additional hour before washing with PBS and then adding cell media suspension.
One important note - Your coverglass must be high quality (many are partial to German glass).  You must in any case first strip all oils and debris off your coverglass before coating to achieve good cell adhesion.  You can use strong acid (>5N HCl) or strong Base >6N NaOH) for at least 1hour.  If you can find some blue plastic coverslip racks that Corning makes then those work well for evenly cleaning several at a time (You can even try buying these racks with coated coverslips the first time and then after they are gone, use the racks to wash/coat your own.  Home-coating is far cheaper and in my experience works more consistently.  For product page, see:  http://catalog2.corning.com/LifeSciences/en-US/search/Search.aspx?searchfor=biocoat%20slides).  
After wash, carefully transfer acid/base waste to appropriate container.  Wash, wash, wash with water until acid or base has been flooded away, using ddH20 on final washes.  Wash in ethanol to sterilize and then once dry proceed with coating above.
Another important note:  It helps a lot to include serum into your plating media (just at the beginning).  This will also prevent clustering, because the neurons will have a healthier jump start to their growth.  You can add it at 25% to your growth media to plate your cell suspension on the coverglass (again pipette as a surface tension dome on the glass).  Incubate for two hours and after cells have adhered, rinse coverglass gently with PBS to remove serum, and then gently add non-serum growth media to the coverglass in well as normal.  Every three days, replace 1/2 volume with pre-warmed growth media. Good Luck!
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i have to find the problems which ouucrs in calcium imaging of neurons? 
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Hi Aashish,
As Pieter mentioned, more information on the specific setup and issues that you are having would allow the community to give you more targeted responses. That said, one resource that I have commonly made use of when having issues with calcium imaging in a cell culture context is a methods paper from Amy Palmer when she worked with Roger Tsien (RIP).
I've attached a link. Best of luck with your experiments.
Joey
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We carried out a WB in nuclear extract of mouse striatum 2h after a stimulus. CREB is detected at 43 KDa, but p-CREB seems to appear at 70 KDa. Has anyone the same problem? Could be a complex of p-CREB with another protein as CBP?
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Very likely that it is not pCREB, but some other protein that just happens to have an epitope similar enough. Most antibodies against phosphorylated residues, even the very best monoclonals, typically have more than 1 band clearly visible. 
Have you BLASTed the sequence that the antibody detects? Might give you more clues. However, most of the time, the antibody manufacturers don't give too much details, so in the end all is just speculation.
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Is it possible to chemically link a small peptide to the side chain of a Heparin sulfate proteoglycan? I know there are antibodies available, but I am specifically looking for a small peptide with less than 10 amino acids.
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It probably is possible, but good luck documenting it. The Heparin sulfate molecule is an extended mess of ill-defined and ill-ordered sulfo-glycans strung on a protein core. Its best use is as a bulk molecule for taking up space. Son't waste your time getting into its fine structure - there really isn't any. Just attempt you linkage, and then see if the product has immune properties that are helpful to you.
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Dear Fellows,
I am trying to induce LTP in horizontal hippocampal slices (400 um) from ~8 month old mouse, stimulating CA3 shaffer collaterals and record from dentrides of CA1 pyramidal neuron. 
After a few rounds of experiments, I found that the theta stimulation protocol I am using now is not sufficient to induce LTP in every slices from WT animals. Stimulation strength was set to provide fEPSPs with an amplitude of ∼30% of the maximum (stimulation pulse duration 0.1 ms). I collected the baseline every 30 s for 30 min. Then, LTP was induced using a theta burst protocol (TBS) consisting of 10 bursts at 5 Hz, and each burst consisting of four pulses at 100 Hz, with a pulse duration of 0.1 ms. TBS was repeated for 3 times, with 20 s interval. For the slices on which TBS with 0.1 ms pulse duration failed to induce LTP, I tried to increase Pulse duration to 1 ms. To my surprise, the same TBS protocal with 1 ms pulse duration can induce LTP on every slices I have tested. 
So my question is as in the title, can I use 1 ms pulse duration for TBS and 0.1 ms pulse for baseline collection before and after stimulation?  Everything else for stimulation pulse remains the same. I did see some people increase the pulse duration in their publications, like increase the duration from 0.1 ms to 0.2 ms. However, there are some worries and also I curious about why 0.1 ms pulse TBS failed to produce LTP in my slices.
Thank you.
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Some interesting points and good tips from the others here. I thought I'd add that there is quite a long history of different labs doubling the pulse width during the induction phase of LTP for short intervals. Usually this i done for either Theta-patterned stimulation of short bursts of high frequency stimuli (i.e. 0.5 sec of 100Hz pulses). In either case, a 0.2 ms pulse for this brief duration is acceptable if you're using a bipolar electrode. Yes you will stimulate more fibers during the induction, but this just helps to create an associative condition where the post synaptic cell is depolarized more during afferent input, enhancing LTP in the fibers you stimulated with lower intensity stimuli. If you do an I/O curve, you should get a leftward shift across stimulus intensities.
Most labs also start off by working at 50% of the maximal response size for a given area. There may be small differences in dorsal/ventral LTP with some paradigms in the CA1 region, but you should keep track of where you are in the hippocampus anyways. My students always keep track of the slice number and usually even quickly draw the shape of the hippocampus. With transverse sections, it becomes quite elongated the more ventral you go. 
Positive controls for LTP are tricky. Really a positive control shows you should get LTP when you expect to, so if you start the day and end the day with slices you can get LTP in, then you're showing it's possible. You could also have an antagonist in place and block LTP initially, and then show you can get LTP when it's washed out. Neither are satisfying positive controls, and your paired-pulse tests and an I/O curve pre and post for every slice is a better way to show you have good slice health for the duration of your recording. 
I hope this helps. We have some reliable ACSF mixtures and stimulus procedures for mice described in the following papers. 
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I am targeting the serotonin transporter in rat striatal tissue. I have done this successfully using whole cell prep. I am now using a synaptosomal prep and instead of my bands coming out around 65-70 kDa, I am seeing a band coming out around 50 kDa that looks like the one I want, but the ladder hasn't shifted. Granted I am using an old stock of primary as well. Is there some factor that would cause a drift in bands?
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Hi Melissa,
unlikely that it is the fault of the old stock of Ab, unless it was badly stocked and is somehow degraded so that it recognizes something different, but then it should be the same for your tissue extract... Just in case you want to make sure, you should have the tissue extract next to the synaptosomal extract in the same WB...
Typically when you obtain a smaller band than expected in WBing, it is protein degradation. I'm not sure what protocol you use for synaptosomal preparation and what for the lysis, but it is possible that in the process you suffered protein degradation. We put the greatest effort in the tissue dissection, so the animal is very quickly sacrificed by decapitation, the head immediately placed on ice, the dissection performed in a few minutes on ice (or a metal slab placed on ice) and the tissue quickly homogenized. Then the prep is performed always using ice-cold buffers, refrigerated centrifuges and the cold room. Finally, the quickest and most efficient lysis for synaptosomes if you only need to run a WB is with 2x SDS protein lysis buffer (Laemmli or similar).
Last point, it is also possible that you don't have enough Sert in striatal synaptosomes to detect it by WBing. I'm not completely sure, but as far as I know 5-HT release is not particularly synaptic, there is high bulk or volume release from the axonal shafts and quite a lot of transporter will be located all along the axons. If you isolate synaptosomes I don't think you get so many from serotoninergic projections, most synaptosomes in striatum are from the cortico-striatal glutamatergic projections (these are essentially the spines of the medium spiny neurons). You might know better...
Anyway, are you exposing a similar time as the WB with whole tissue lysates? And how much synaptosomal lysate are you loading? If this signal at 50 kDa is coming up after a long exposure it could be likely non specific. You might try loading more synaptosomal protein, up to 25-30 ug (a real lot for synaptosomes, of course, but just to see if something specific comes up...). If the signal is very sharp it is difficult to think at degradation, unless there is some sensitive site where the protein can be cut (a PEST or similar site). Normally there is some background smear when you have degradation. Do you only get this band at 50 kDa or are there other non specific bands, less intense?
Good luck.
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We are interested in looking at vascularization and angiogenesis in some human brain samples. The ideal way would be to use immunostaining but our lab is not developed for IHC or any immunolabeling experiments. So I was wondering what would be a good marker to look at vascularization using qPCR? Any suggestions are welcome, and it would be best if you can also provide some citations. Thanks!
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Thank you for the suggestions!
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How is the performance? As compared to virus? My plan is to deliver oligos in cortex 
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Dear Yu Wang,
I would like to offer you the possibility to try our reagents as an alternative for your application. Two strategies may be used:
either Magnetofection: our in vivo line of magnetic reagents are designed for in vivo targeted transfection and transduction. This combines magnetic nanoparticles and nucleic acid vectors that are retained after injection at the magnetically targeted site. In this way, systemic distribution is minimized and toxicity is reduced.
or "classic" transfection reagents that are really efficient in vivo.
Results have been obtained in Rat brain for DNA transfection using the methods presenting above.
Would you like to try them, please do not hesitate to contact me via ResearchGate or directly at tech@ozbiosciences.com;.
best regards,
Cedric
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Dear Community,
this is about Action Potential transduction speed over a Neuron!
Reaching a Neuron, an AP will first go through the dendrites, then reach the cell-body. Afterwards, it will go over to the Axon, then find a Synapse to give the AP to the next Neuron.
I need speed-estimates of every single one.
I found some estimates for conduction velocity to be about 1m/s for unmyelinated Axons.
The transduction over dendrites and cell-body might be in the same range!?
 The transduction over a synapse is fast, but I am interested in a cell-cluster, so I have about 10 Synapses.
It would help us a lot!
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"Reaching a Neuron, an AP will first go through the dendrites, then reach the cell-body. Afterwards, it will go over to the Axon, then find a Synapse to give the AP to the next Neuron."
This is not at all how action potential firing unfolds.
Graded postsynaptic potentials will travel from the synapses down the dendrites and integrate at the soma (and in the dendrites). If the sum of these reach a certain voltage threshold at the axon initial segment (located close to the soma), an all-or-nothing AP will propagate out the axon from here. The AP will also back-propagate through the dendritic three.
As Nikolaos correctly writes above, the AP does not cross the synaptic cleft, instead here the signal is neurotransmitters that diffuse the short distance to the postsynapse, where they give rise to graded potentials, and so forth.
As for your model, each neuron commonly have thousands of synapses, so you probably have to describe better what you mean by a cell-cluster with 10 synapses.
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I want estimate different neurotransmitter levels in the rat. For which I prepared whole brain tissue homogenized  in 0.1N perchloric acid with Sodium bisulphate.
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Dear Atif,
It depends on what you are trying to find out. Basically each neurotransmitter exists in independent neurons/neuronal clusters/terminals/ and also at some 'nuclei or in a single neurons co-existence of neurotransmitters are present. By analyzing a specific area in a brain circuit may provide you the most valid and meaningful information, because the area could be a known cluster of neurons or terminals for a particular neurotransmitter where it is synthesized or getting released, etc. and there could be modulator-neurotransmitters/neuromodulators present in these areas. Since much information is available about specific neuronal loops and connections, it is desirable to get information from such areas of the brain where it matters or in other words has specific functions. A typical example is nigrostriatal loop of dopaminergic connections and the neurotransmitter at the striatum could provide you much information (eg. on motor activity), than from a whole brain homogenate. Similarly assaying dopamine at mesolimbic pathway provides information on emotion and probably cogntion! These explain why I stated 'whole brain homogenate may give not much information'. 
All the best.
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Im planing to suppress neurons excitability by over-expressing the Kir2.1 channel. My question is, besides electro-physiology is there any other method which I can use to validate this method. As in a method that will show that the adenociral vector injected overexpressed Kir2.1 and suppressed the neuronal excitability. 
Thank you. 
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Hi Nanthini,
Indeed, you can measure a number of electrophysiological parameters to assess the decreased neuronal excitability upon expression of Kir2.1. It has been done in previous studies numerous times. For instance you can measure spontaneous / induced firing, resting membrane potential, input resistance, etc... if you do not have access to electrophysiology, you may want to use voltage-sensitive dye and field stimulation to assess some excitability properties of your neurons. Also calcium imaging properly used may provide you with some information. If you are expressing Kir2.1 in vivo, you can certainly look at behavioral properties depending where you are expressing your channel. And of cause, you want to make sure that all of this is indeed mediated by manupulating Kir2.1 expression that you can validate by western blot, immunostaining, or patch clamp.
Hope this helps. All the best. Norbert
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I am using 4-OH-Tamoxifen for intraperitonial injections in mice at 50mg/kg. I used to follow a method where 4-OH-tamoxifen is dissolevd at 20mg/ml in ethanol and then oil is added to get final concentration as 10mg/ml. The ethanol is then evaporated in centrivap. However I think this gives me variation. Is there a better protocol independent of sonication and centrivap? 
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Do you use the vortex after add the oil ?  Mix well the oil and ethanol before to use the centrivap  would help.
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Hi, Is it possible to structurally modifiy rat brain hippocampus for the accumulation of BDNF and Glutamate neurotransmitters for enhancing memory function?
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I want to explore the mechanism of hippocampal LTP and the role of BDNF in synapse. So, is there any anatomical method for remodelling of the hippocampus region? 
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Which amplifier is best for in vivo recording, especially for very weak signal?Which amplifier can be used for recording identified neurons and nerve fiber firing?
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Dear Yan, before one can answer your question it is required to clear what exactly you want to record. For example do you plan intra- or extracellular recordings. Do you plan to use glass pipette filled with electrolyte which is placed inside a single nerve cell for intracellular recording or do you plan to use metal electrodes for extracellular recordings. Depending on the answer on this question you have to choose a preamplifier that is adapted to the electrode of choice. Micropipettes and metal microelectrodes usually require different amplifiers. The input characteristics of these preamplifier have to be adapted to the recording sensor. For example the input impedance of the preamplifier should be 100times higher as the microelectrode impedance in order to measure the full amplitude of the biosignal. Further requirements for the preamplifier are low leakage current of the input and a good response time. Metal electrodes used for extracellular recordings usually have lower impedance values as micropipettes and therefore the requirements for a preamplifier are not so high as for a micropipette preamplifier. The preamplifier is followed by a main amplifier. Depending on the signals you want to record you can choose a main amplifier/filter system. For example if you plan to record single unit activity it is recommended to use a bandpass filter (e.g. 500Hz to 5kHz). So there are much more options depending on the recording application. To give you an optimal recommendation it is required to know more details about what you exactly want to measure and which components (e.g. electrodes pipettes) you already have in your lab. Best wishes, Dirk
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Hi, I am looking at using PuraMatrix in 3D cell using iPSC derived neurons. Does anyone have a protocol I could follow and any tips/advice for a successful culture? Initially I would like to carry out some IF to look at neuronal markers, so a thin layer 3D culture. Many thanks, Alys
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Maybe you can take a look at this paper?
Best regards from Germany
Vladimir
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Hey everyone,
I know the question seems paradoxical, but I am currently trying to measure sEPSC in the medial Hippocampus CA1 cells in adult mice voltage clamped at -70mV, and I find that there are these huge (1500~4000 pA) currents that pop up every minute or so (image attached). The internal solution is CsMeSO4 based and contains 5mM QX-314 (alomone labs). From my understanding, the voltage clamp and QX-314 should both stop the clamped cell from firing, but it does. Frustrated that I was, using the same int sol I changed to current clamp and found that there are depolarizations that go up to 10mV, which is why I am saying that the cells are "firing." Has anyone experienced this? As a last resort I borrowed some internal solution with the same composition from another lab, but got the same results. The alomone lab website, by the way, says that the particular chemical, at 5mM the Na current is reduced by ~20%, and complete blockage of sodium currents require 50+mM concentrations. But from what I know the conventional knowledge is that 5mM is enough to block them, so I am at a complete loss as to what to do. Any help would be appreciated. Thank you all in advance!
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Dear Haram,
The neurons you are trying clamp are very large and therefore there is a problem with space-clamp issue, meaning that you are not actually fully clamping the cell at the holding voltage outside the immediate location of the electrode tip. Action potentials are coming from the axon hilux are sometimes difficult to clamp, especially if you are using small tip (high resistance) glass micropipettes. What is your access resistance readings? My recommendation would be to use larger tip electrodes 2-3 megaohm (~1 micron) in the bath solution. The larger the electrode-membrane interface the lower the access resistance will be, meaning you will be able pass larger amounts of current that will give you better voltage-clamp. The access resistance ideally should be less than 10 megaohm, especially in neurons with larger dendritic trees. This should also help passage of QX-314 to the intracellular compartments easier. For inspiration see my following publications.
Best wishes, Refik
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When I using NeuN antibody(MAB377 | Anti-NeuN Antibody, clone A60), I found that the signal was almost confined to nucleus. While applying to cultured hippocampus neurons at DIV 10, I found the signal not only in nucleus but also in cytopasm, I could even find strong shaped distribution signal in nervous process. Can anyone tell me WHY? Thanks.
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iF  you stain adult tissue with NeuN and use ABC peroxidase methods with perfusion-fixed tissue processed freely floating, you will see high specificity for neurons and no staining of glia, but if conditions are optimized, the staining will be in nuclei and in the dendrites of many neurons (usually nuclei stain stronger than other structures.  If you used young animals or used young animals for cultures, the staining is more likely in nuclei.  What this means is that proteins reacting with the antibody are higher in adults than in young animals and expression is likely low in cultured tissue compared with intact sections.   From your photo, I assume you are staining with a fluorophore-tagged secondary antibody. You should know that the antibodies against NeuN will stain optimally at 1:30,000 if tissue is well fixed and incubated according to our recommended ABC protocol (Hoffman, G.E., Sita L.V. and Le, WW. Importance of titrating antibodies. Current Protocols in Neuroscience Chapter 2 Unit 2.12, 2008. PMID: 18972376).  When you are comparing staining of sections with cultures, the optimal concentration of the antibody will likely differ depending on how well the tissue is fixed and how long you incubate the tissue with the primary antibody.  You could find that by optimizing the concentration of primary and using a more sensitive technique both conditions might show similar localizations. This is what NeuN looks like in a spinal cord section optimally stained (1:50,000, 48 hrs with primary, and rest of protocol according to our standard protocol in the paper indicated above).  I see the same with optimized fluorescence but primary concentrations can be 50x higher if using a direct fluorophore secondary instead of ABC.
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Hi all,
I have someone's student starting to measure NMDAR mEPSCs from cultured neurons and I instructed him how to proceed from my experience in brain slices. He, however, went on to find various options for his solutions and settled for a low magnesium extracellular with a sort of standard internal of CsMeSO3 + lidocaine. 0.5 mM TTX, 10 uM CNQX and 10 uM biccuculine are added to the bath. When he goes to +40 mV to relieve the Mg2+ block of NMDARs the membrane resistance decreases from 300 MOhm to about 100 MOhm and the recording becomes unstable to the point of being unusable, with the baseline going up and down as it pleases. It looks much better at -70, although still "messy". This is likely due to some conductance being activated under these particular conditions and I'm leaning towards the low extracellular magnesium as the culprit. The problem is they are trying to replicate someone else's conditions so I'm at a loss as to why this is not working here... Never had this sort of issue in slices at +40 mV in normal Mg2+ ... recordings did seem to get noisier but were well usable. Anyone experienced something like that?
Regards, 
Paulo.
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Paulo,
have you tried to use QX-314 instead lidocaine in your internal solution? You have the same problem without lidocaine? Have you tried?
Wishes
Luca
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I'm running experiments and I need to silence the mouse DRG in culture. I'm thinking of either Lidocaine or Toxin cocktail (TTX with conotoxins). Does any one have experience in this and willing to share some information regrading protocol conc. duration etc.
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Sorry I just realized that you are doing it on cultured neurons. Why not in vivo? I would just use TTX in that case around 1-10 micromolar should block action potentials. See my 2005 JPET electrophysiology paper on cultured DRG neurons. Another drug you may consider is riluzole, an FDA approved drug, though it does not block all action potentials (see my 2013 NeuroMethods paper).
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In a network of coupled oscillators (diffusive coupling) while it is intuitive to expect that the magnitude of coupling increases with increase in concentration of coupling factor (provided the diffusion rates and other factors are held constant)....
1. Would this increase be linear, sigmoid or does the coupling strength follow an inverted U shaped curve with an increase initially and eventual drop?
2. Does higher concentration of coupling factor lead to unstable network, or in other words will the network attain local stability but is easily susceptible to mutual desynchronization by an external agent? 
3. Also, is this trend different for mean field coupling?
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Fascinating questions. Not sure I can answer them all directly but I can recommend some reading if you are interested in the topic. It turns out there are many people within the circadian field who are interested in questions such as these and have used mathematical models of the circadian system to try and answer them.
1) I would say that it is sigmoid, at least based on mathematical models that test increased coupling strength. This should hold true for your question If you assume that increasing the concentration of a diffusible factor indeed increases coupling between the oscillators (up to the saturation limit of the receptors for that signalling molecule). I am not aware of a scenario where it would be an inverted U, unless there was some weird biphasic response to the molecule itself. Say if there were two receptors, one with high affinity which increased coupling and one with low affinity that decreased or blocked it. I am unaware of such a system for neuropeptide signalling but that doesn't mean one doesn't exist.
2) Theoretically, increased coupling always leads to greater stability and synchonization of the oscillators. However it all depends on zeitgeber strength. If the coupling agent is a weaker zeitgeber and some other external factor is a stronger one, then it is likely that the entire system would be more easily shifted by the external factor. In the case of strong coupling between oscillators, all oscillators should still remain synchronized to each other even if they are perturbed by an outside force, i.e. they would all move together or not at all.The opposite would be true for a weakly coupled system where a strong outside force on some (but not all) cells could desynchronize the system. I think it is safe to say that, in general, increased coupling will always convert local stability into global stability (assuming all cells are equally coupled and are equally perturbed by outside forces of course). The situation changes very quickly and  dynamically if you have multiple diffusible synchronizing factors released from different subpopulations within the network. This is much more like the case of the suprachiasmatic nucleus, with multiple diffusible factors all released from spatially distinct subpopulations of cells with both local and long-range connectivity (think small-world models). In general though, this more complicated scheme allows for greater stability and synchronization across the network while still allowing phase-resetting stimuli to entrain the system in a predictable and controlled way.
Abraham, U., Granada, A. E., Westermark, P. O., Heine, M., Kramer, A., & Herzel, H. (2010). Coupling governs entrainment range of circadian clocks. Mol Syst Biol, 6.
Achermann, P., & Kunz, H. (1999). Modeling Circadian Rhythm Generation in the Suprachiasmatic Nucleus with Locally Coupled Self-Sustained Oscillators: Phase Shifts and Phase Response Curves. Journal of Biological Rhythms, 14(6), 460-468.
Ananthasubramaniam, B., Herzog, E. D., & Herzel, H. (2014). Timing of Neuropeptide Coupling Determines Synchrony and Entrainment in the Mammalian Circadian Clock. PLoS Comput Biol, 10(4), e1003565.
Bordyugov, G., Granada, A. E., & Herzel, H. (2011). How coupling determines the entrainment of circadian clocks. The European Physical Journal B, 82(3-4), 227-234.
DeWoskin, D., Geng, W., Stinchcombe, A. R., & Forger, D. B. (2014). It is not the parts, but how they interact that determines the behaviour of circadian rhythms across scales and organisms. Interface Focus, 4(3), 20130076.
Kori, H., Kawamura, Y., & Masuda, N. (2012). Structure of cell networks critically determines oscillation regularity. Journal of Theoretical Biology, 297, 61-72.
Vasalou, C., Herzog, E. D., & Henson, M. A. (2009). Small-World Network Models of Intercellular Coupling Predict Enhanced Synchronization in the Suprachiasmatic Nucleus. Journal of Biological Rhythms, 24(3), 243-254.
Gonze, D., Bernard, S., Waltermann, C., Kramer, A., & Herzel, H. (2005). Spontaneous synchronization of coupled circadian oscillators. Biophys J, 89, 120 - 129.
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Hi all, I've been facing some problems with microglia in my secondary astrocyte cultures. I use swiss mice(p0-p2) and, although there are quite few in the primary culture (in culture flask), after the passage, which is around 6-7DIV, they start to proliferate and assume an activated/amoeboid morphology (in 6 well culture cluster). If I plate the primary culture in 6 well clusters, to avoid the subculture process, they also appear and already 'activated/amoeboid'. If you guys have any suggestion, please let me know. Thanks!
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Matheus, I am not certain what you mean by "secondary" culture. Nevertheless, you should be able to sufficiently remove microglia from your astrocyte cultures so that you achieve a reduction in microglia to the point that they represent less than 0.5% of your total cells. I have used anti-CD11b MACS beads and consistently achieved high astrocyte purity. I have attached an article where the procedure is briefly described. Results are presented in Fig.6A and 6B.
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Brain
Frozen at -80 degrees since 2011 (no reason to believe samples were tampered with)
activated PPARy ELISA 
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Hello..Its very difficult to get result of such a long time stored sample. But I suggest you may try, again centrifuge the sample aliquot and then perform the ELISA. May be it will work.
Gud luck..
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To find the therapeutic application of the peptide
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try SATPdb to find therapeutic application of peptides
SATPdb: a database of structurally annotated therepeutic peptides.
paper link
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Does anyone know any mild model to induce neuroinflammation different from LPS model in rats?
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Also consider the type of immune reaction you are after (e.g. involvement of specific cells and acute inflammation versus chronic/prolonged). I have found that microglia react very quickly and easily to single cytokines or other toxins, while astrocytes often require a cytokine cocktail (e.g. TNF-alpha + IL-1beta). If you are after a chronic inflammation, then initial microglial activation can also lead to astroctye activation over time. 
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At the moment I am trying to isolate Purkinje cells from mouse cerebellum. I have a protocol which contains steps like 3% PFA perfusion, homogenization of the cerebellum with douncer, filtering through 100um pored nylon filters and stainings for FACS sorting. Main buffers are HBSS and PBS.
After the sorting (I sorted over Parvalbumin, although I would have GFP expressing Purkinje cells) I extracted RNA from my cells of interest by usin the RNA Kit of Macherey-Nagel. I got in total 198ng RNA with a RIN of 3.1 and bad purity. The extraction of the RNA happens 2 days after the perfusion. Until then I stored the homogenate at 4°C.
Now I am trying to shorten and improve the protocol. Does anybody has any experience with isolation of specific cell type from brain followed by extraction of RNA? I will need the RNA for RNA-Seq to compare wildtype with a knockout.
Thanks for any suggestions.
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Have you tried to sort them directly into triazol buffer in 1.5 ml size microtube; then, right away do the standard triazol-chloroform RNA extraction?. It did work for mouse neuro progenitor cells (around 30-50 thousands cells) for RNA-seq.
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We want to patch clamp cells with GABA A receptors, without the need to transfect the cells first. A colleague told me that N2A cells express GABA receptors without the need for transfection, but I'm having trouble finding papers that confirm this. Do the cells need to be differentiated to neurons before they'll express GABA A receptors, or do they already express these as primary cultures?
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Dear Professor, We studied also early neural development in the avian embryo. We discovered that early neural development (pre-neurulation) is provoked by induction by the endophyll (primary hypoblast).Indeed endophyll orients and organizes the early head region (Callebaut et al.,1999: Eur. J. Morphol. 37 : 37-52. On the other hand by placing a quail Hensen's node (Stage 4-5 V) on the deep side of the upper layer of an isolated anti-sickle region from unincubated chicken blastoderms, we observed the appearance of a quail floor plate, a quail chorda and also quaill cells in the median roof of the definitive endoderm. ( Early steps in neural development : Callebaut et al.,2006: J. Morphol. 265 : 793-802. See our E-book : From ooplasms to embryo : The "embryo in box" model (Edit. acco, Leuven, Belgium).With kind regards.
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Given Fabbri et al. (2014), has anyone created a simultaneous computational neurogenetics analyses
of GWD-CGA-fMRI public accessible data to elucidate neural substrates (e.g., Miller et al., 2015) to various DSM-5 Axis I disorders to inform etiology of developmental psychopathology to inspire innovative epigenetic studies to prevent development of psychiatric disorders [(e.g., reversing an underlying neural substrate to behavioural inhibition (Bellgowan et al., 2015) via a proactive disinhibition intervention (e.g., play therapy) to prevent development of internalizing disorders in children)]?
Bellgowan et al. (2015). A neural substrate for behavioral inhibition in the risk for major depressive disorder. Journal of the American Academy of Child and Adolescent Psychiatry. DOI: 10.1016/j.jaac.2015.08.001.
Fabbri et al. (2014). From pharmacogenetics to pharmacogenomics: the way toward the personalisation of antidepressant treatment. Canadian Journal of Psychiatry. PMID: 24881125.
Miller et al. (2015). Meta-analysis of functional neuroimaging of major depressive disorder in youth. JAMA Psychiatry. DOI: 10.1001/jamapsychiatry.2015.1376.
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Now comes an innovative pre- and post-fMRI analytical technique at an individual level of analysis-functional connectome fingerprinting (Finn et al., 2015) which may document the efficacy of a treatment intervention to reverse an underlying neural substrate to behavioural inhibition (Bellgowan et al., 2015) to prevent development of an internalising disorder. That conceptual model can be applied to other psychiatric disorders as well.
Finn et al., (2015). Functional connectome fingerprinting: identifying individuals using patterns of brain connectivity. Nat. Neurosci. Oct. 12. doi: 10.1038/nn.4135.
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which serotype of AAV-shRNA is best for rat mid-brain neurons? Thanks.
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AAV DJ serotype is also good. 
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Hi! Has anybody come across the same problem? We are doing MRI experiments on the ex-vivo brains. Unfortunately all the brains that I have extracted so far have air bubbles inside :( It is air for sure - it causes artifacts in the T2*-weighted images.
I tried leaving the brain in a skull ( just removed the muscles and skin, but did not extract the brain), and it is still the same! I do perfusion in a standard way: PBS/PFA perfusion, cutting the head with the guillotine, careful brain extraction, yet here are those bubbles.. 
Today I was double-checking there are no bubbles in the perfusion lines - it is almost certainly not the cause.. May be the guillotine? Or something else?.. I do it the same way as people in the institute do, but nobody yet has looked at the MRI images, so perhaps it is just an MRI-specific issue..
I have just finished my experiment, and I have 30 rats to perfuse, and I am really worried they will have air bubbles and won't be usable... Any suggestions would be most welcome..
cheers!
Katya
p.s. attached is the image of the brain left in the skull (imaged at 1T).. 
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Hi Ekaterina,
For the move from PBS to Florinert - density is your friend. Fluroinert is 1.6 times heavier than water.  You can place the water covered brain in PBS and push it down into the Fluroinert.  I use the least viscous fluid I can find to avoid pockets of PBS or Air in the brain.  If you can get the version that has a viscosity between 20-100 centiStokes it is a great help. 
As for the extraction time its not that much of a difference than extracting in air - less than 10 minutes at most depending on your set of instruments.
Once the brain is extracted I've not usually taken much trouble in moving from PBS to Fluorinert.  You will find that as you experiment with your technique there will be a procedure that works best for you.  You can easily check for any small air pockets by running a long TE gradient recalled echo MRI scan.  This is very sensitive to local magnetic field variation which the air pockets great.  If any are found you can remove them by manipulating the sample under the Fluorinert.  This can be done by using tapping agains a hard surface, a vibrating surface or attaching a water aspirated vacuum while manipulating the sample.  Fluorinert is a high-temperature pump oil so it will have no appreciable vapor pressure under a soft vacuum.
I hope this helps.
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I want to do a double labeling Immuno-histochemistry co-localization experiment for c-fos and TH in free floating rat brain sections. both the primary antibodies are raised in rabbit. Apart from using Alexa-555 tagged anti-c-fos secondary Antibody , which fluorophore would be best suited for another secondary antibody which reduces overlap or bleed-through load???
Most of the fluorophores either have high bleed-through or do more overlap in their emission spectra or have an emission range which in co-localization experiments makes it difficult to visualize.
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You can directly conjugate the fluorophores to primary antibodies if you have to use the antibodies from same species.  With appropriate filters, Cy3 or Alexa 555 and Cy5 should not have any bleed-through.
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I have had this ICC working for several years and have used it for multiple experiments, but recently it has stopped working. I am now having nearly every cell in the brain labeled in every trial I do. I initially tried ordering a new lot of the primary antibody, but that made no difference. I have modified the concentration of the secondary and the time in the DAB and neither of those helped. I tried doing a no primary control to rule out nonspecific binding of the secondary and a no primary or secondary control to rule out ABC binding to endogenous biotin in the tissue. Both of those trials came out clean, with no staining, suggesting that the problem is not with the secondary or the ABC. We have tried replacing the paraformaldehyde (our brains are not perfused, so fixation is the first step of our IHC), the methanol, the H2O2, the normal goat serum, and the DAB tablets and none of these things have solved the problem. We also thought that contamination in the water may have been the problem, but I got the same result when using all solutions made with HPLC grade bottled water. Finally we thought there may have been a problem with the tissue since it had been in the -80 freezer for 2+ years and may have experienced temperature variation, but I recently included sections from a brain that was collected and sliced within the previous week and that did not solve the problem.  I have run out of ideas for what could be causing the excess staining.  Does anyone have suggestions on what I could try?
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The steps you are following to identify the culprit are all right. Write down a flow chart of the problems. Some suggestions for it: Are you getting the same high background when using other atbs? If NO, follow recommendation from Srinivasan, above. If YES, go to a colleague who is now running (successful) IHC and repeat your experiment in his/her setting. If GOOD results there, start replacing reagents, one by one, etc.Good luck
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We know that melanopsin is sensitive to blue light with a peak around 480 nm and that melatonin suppression is most sensitive to blue light with a peak around 460 nm. Furthermore, it seems that melanopsin is important but not essential to circadian photoentrainment. I have found some older reports showing that circadian phase shifts are more sensitive to short wavelength light.
Does anyone have any idea of how important blue light is to circadian photoentrainment, preferably based on recent research?
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Lots of experimental evidence are available to support that monochromatic light shift the clock, especially blue light compared to red light. Blue light shifts the clock (circadian locomotor activity rhythm)  but not red light. When we keep animals in constant DD, people have been using constant red light.  if you keep the animal in DD, blue light exposure definitely shifts the clock. 
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LPS (1ug/mL)-stimulated microglia are positive for CD68, but so are Vehicle (serum-free medium) treated microglia. I used fluorescent microscopy to visualise them. Fluorescence intensity for CD68 in both groups show no significant difference! So I'm skeptical about using anti-CD68 antibodies to identify activated microglia. Please suggest where I could have gone wrong.
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There is no single test to identify "microglial activation", because there is not uniform state called activation. Microglia have different activation phenotypes, which are linked partially conflicting functional features (Dr. Carol Colton's has couple of enlightening reviews about this). Different stimuli induce different responses and surface protein expression might not always reflect on those. E.g. LPS typically will induce NO release, some pro-inflammatory cytokines and very distinguish morphological transformation. I am not sure what happened to anti-inflammatory cytokines or trophic factor release with LPS, as I rarely use it myself. In your situation, LPS did not change CD68 expression, but it might be that CD68 basal expression of your primary(?) microglial cultures was elevated due to serum-deprivation, or simply it just is not sensitive method to detect "microglial activation". My suggestion is that you identify several functional outcomes typical for your stimuli (i.e. LPS) in your culture conditions (i.e. serum deprivation) and then see how those are changed with what ever treatments you plan. 
One more point, in papers people (including myself) use the term "microglial activation" in lack of the better term, but nowadays referees usually expect multiple outcome measures, particularly in vitro, in vivo is still a pickle...
I call microglia the drama queen of the brain, so on that note - Good luck! ;)
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Hi,
I am a new student who is going to do some electroporation on primary cortical neurons. Once harvesting cortical neurons, how much for amplitude and duration for the pulse shall I use? I am using Bio-red GenePulser, with 0.4 cm cuvette. However, I could not found useful protocol from the company for cortical neurons as well as google. Someone only mentioned the name of program they used on their machine. 
So, I want to know, how many neurons and plasmid you guys used, and the detailed protocol for electro-pulse (amplitude, duration, number of pulses etc.).
Thank you, and wish you a good day!
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Chenglong,
Sorry for the delay in my reply. I've been busy with experiments and paper.
To answer your questions in the order they were asked: 1) if your neurons survive the harsh conditions of trituration and electroporation, they will start to send out neurites in 24 hours if not overnight. No dying or dead neuron will send out processes, so I would call those survivors and yes they will grow in size and neurite complexity. We typically see GFP expression from electroporated vectors in 24 hrs. However, how long your fluorescence persists depends on your plasmid vector and half life of the fluorescent protein. We use CAG driven plasmids for life-long expression of fluorescent protein. In our hands, CMV driven expression doesn't last very long, a couple of days at best. If you want to see spine, DIV 14-21 should be long enough for your cultured neurons to send them out and long enough for some to mature into stubby or mushroom shaped ones, if you have a healthy culturing system. 2) Since your neurons don't divide, the number of plasmid in your transfected neurons should not change, in theory. However, whether and for how long the expression persists depends on the promoter used in constructing your plasmid. For indefinite continuous expression, I recommend CAG promoter not CMV.
best of luck,
L
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We are having trouble preparing a solution of 7,8 DHF to administer in drinking water to our mice.  We are preparing the compound in DMSO and sucrose solution but have great difficulty getting the DHF into solution and subsequently keeping it in solution.  Has anybody any hands on experience in making up this compound?
Thanks in advance,
Sinéad
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Hi Nick,
Yes we still kept the sucrose and DMSO in there but just the more basic pH of the water made the DHF go into solution much easier.
Sinéad
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I was having a discussion about the nature of phosphenes and it was proposed that the generation of these light like percepts could be an effect of the tms pulse being conducted down the retinal tracts, stimulating the retina, and then being passed back to the cortex for further processing.
This seems like a very slow process, and a number of  sources would agree with me that this does not sound like the most likely mechanism.  However, does anybody know of any recent (last 10/20 years) research that has investigated this?
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