NetNotes


imaging chamber off and clean everything thoroughly, then oil it and check again. You can sometimes remove a bubble with the edge of a lens tissue but more oſten that just makes things worse. Keep in mind that it doesn’t have to be a bubble - sometimes these systems just lose their lock and never get it back. For example, if you don’t quite have enough oil then you can easily break the focus lock partway through an experiment. Also, note that if your system has the option to choose between continuous and point-by-point mode, continuous mode is a lot more likely to scotch the entire experiment if it momentarily loses a lock. For this reason, I generally use point-by-point mode for long- term multipoint time lapse experiments, and continuous mode for higher-speed and short-term acquisitions. Another non-bubble-related problem has to do with how far the stage


moves between points. Most systems default their stage to move very fast between multi-points to save time but moving fast over a long distance can break autofocus lock. Tink about this when you set up your multi-points - try to minimize how far the stage moves from point to point and keep an eye on how far it must move between the last first points. If necessary, create ‘dummy’ positions between distant points so that the stage can pause and re-calibrate. Nikon (no commercial interest) has a great feature that automatically re-orders the multi-points to minimize stage travel; with other scope makers you just have to keep all this in mind while setting up the experiment. Slowing down the stage movement during multi-point helps a lot, especially with the continuous focus setting. If you are using an infrared beam autofocus (Nikon, Leica, etc.)


then I don’t recommend 1.33 RI water-matched oil. Tose autofocus implementations need a refractive index mismatch at the coverslip, so they oſten struggle to get and hold a lock with water or water- mimicking oil immersion. I agree with others that it’s a good idea to thoroughly spread the oil around before starting the experiment. Also, use an objective heating collar and give the whole system an hour or two to thermally stabilize before you leave it unattended. Timothy Feinstein tnf8@pitt.edu


In an exact set up as yours, I have encountered similar problems.


To overcome the issues I did the following: 1) Before putting the multi-well chamber on to the stage, I made sure that the microscope was well-equilibrated with the incubation chamber at 37°C. 2) Coat the bottom part of the glass coverslip with a film of oil by rolling the cylinder part of the glass tip applicator. In my experience, using a glass tip applicator reduces bubbles. 3) Put oil on the objective and set the plate on the stage. 4) Make contact between the objective and the plate and bring the cells into focus. 5) Allow the plate to equilibrate at 37°C for 30 minutes. 6) Select all regions of interest in the soſtware and set the perfect focus (reflection-based focus value) for each point. Because of a slight slant, as I move from well to well the focus will change. Not waiting for 30 minutes to equilibrate also results in incorrect reflection- based focus values for the entirety of the experiment. Move the plate through this path manually a couple of times to make sure that oil is evenly spread out and there is no possibility of a bubble forming later due to insufficient oil. Note: Initially, my experiments failed because the thermal driſt with only the stage top incubator was overwhelming for a reflection-based system to manage aſter a few hours. Using both the stage top incubator and an incubator that encloses an entire microscope, I have done time-lapse for up to 20 hours. Gaurav Joshi gnjoshi@emory.edu


Imaging of Organoids Confocal Listserver I have a user who would like to image live organoids via confocal/ multiphoton without jeopardizing the sample. Has anyone figured out


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a material in which the organoid can be placed that would eliminate movement/driſt during imaging, without damaging it so the sample can go back into culture aſterwards? So far, the papers I’ve looked at on this topic cryoembed them at a certain time point for immunohistochemistry. My user’s lab has samples which are expressing GFP and RFP and they would like to image them in vivo. Placing them in a mix of matrigel and culture media would slow them down temporarily, but in my experience, matrigel attenuates the signal. We have a multiphoton system on an inverted Zeiss platform (LSM 710). We can image using an objective inverter and dipping lenses, or from the bottom through a cover glass dish. Since we are working remotely, I was hoping to get some direction from this group.Mary Ellen Pease mary.ellen.pease@gmail.com


We had an interesting talk by Daniel Fulton from the University


of Birmingham, UK (https://www.birmingham.ac.uk/staff/profiles/ inflammation-ageing/fulton-daniel.aspx) where he spoke about long- term live imaging of oligodendrocyte myelination in organotypic brain slice cultures. Maybe some of his techniques might be useful? His talk is at the end of the session located at this link: https://www.youtube.com/ watch?v=0L7tDQ2vDsM Phillipa Timmins phillipa.timmins@aurox.co.uk


Staining of organoids within matrigel is a challenge and results in


bad signal-to-noise ratios, but if the organoids are expressing GFP, or even better long wavelength fluorophores, imaging can be performed within the matrigel matrix. A dilution of matrigel with medium works well and dilutions of up to 1:5 are applicable. To increase the number of organoids, close to the coverslip, you can place the dish on an ice pack while you seed the matrigel-medium-organoid suspensions and keep it on ice for 5 minutes. Within cold matrigel the organoids sink by gravity and you’ll have more organoids within the working distance of the lens. I recommend spinning disc microscopy, due to low photo-toxic stress. Matrigel and 2-photon imaging might be challenging due to the second harmonics of the collagen matrix. Philipp Tripal philipp.tripal@fau.de


Some time ago I was helping a user with their zebrafish embryos.


Te system was an LSM 780 upright with dipping lenses. Te embryos were maintained in position by pressing small dimples into the top surface of (I think) agarose, then resting each embryo in a dimple, covered in water. It was sufficient in most cases to prevent driſt. Michael Doubé mdoube@cityu.edu.hk


Have a look at CyGel. You can get it from Biostatus and Abcam


also distributes it in the US. It’s a liquid at 4°C and it gels around 22- 27°C. It is reversible once you lower the temperature. We have not tried it with 2-photon imaging or organoids, but it worked quite well in our hands with cells and fluorescence microscopy. Zbigniew Mikulski mikulski@lji.org


We kept organoids alive and apparently happy for 12–18 hours


on a microscope stage using embedding in Matrigel. In addition to organoid imaging alone, cells can be added to Matrigel to watch how they interact with organoids. Imaging in the paper below was at low mag, but (not used in the paper below) I fixed and labeled the organoids aſter overnight imaging to see the same ones at high resolution with confocal, so confident this would have worked live at high res too. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5716041/ Michael Cammer michael.cammer@nyulangone.org


We regularly image organoids embedded in matrigel on an LSM710


(invert) for up to 4 days. We’ve imaged GFP, Venus, mBanana, RFP and mCherry, using the standard single photon laser lines. Te organoids have been embedded in a matrigel cylinder around 3 mm in diameter x


www.microscopy-today.com • 2020 November


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