Demo Archive

Room lighting is standard overhead fluorescent, mice are brought straight from the housing room to the testing room right before each trial. Switching to 60-min acclimation plus dim (10-20 lux) lighting made a huge difference for us — WT controls started showing normal center-time exploration instead of near-total thigmotaxis, and our genotype effects became clearly visible again.

To mitigate male cage-mate fighting without resorting to single housing, we implemented a combination of strategies: 1) We preserve odor cues during cage changes by transferring a handful of clean, unsoiled nesting material from the old cage into the clean cage rather than completely washing all odors away. 2) We added crinkle paper and cardboard tunnels to increase environmental complexity, giving submissive mice places to hide. 3) We avoid mixing males from different litters after weaning, keeping sibling cohorts together. This significantly reduced aggression, fighting, and barbering behavior, while maintaining social housing and stable behavioral baselines.

Standard approach that fixed this for us: run 3 training trials per day for 2-3 consecutive days before the actual test day, using the same rod speed/acceleration protocol you'll use for testing. By the end of the training period most mice (WT anyway) should be staying on for well over a minute consistently, and then test-day latency-to-fall numbers actually start reflecting real motor coordination differences between genotypes instead of just "did this mouse ever see a rotarod before." --- Post #4 by rotarod_newbie --- That makes a lot of sense, we jumped straight to testing without any training days. Adding 3 trials/day for 3 days before our real test day.

Also, after each trial, dry mice off promptly and place them under a heating lamp (or on a heating pad, low setting) for a few minutes before returning them to their home cage — this post-test warming step helps a lot with the lethargy/huddling you're seeing afterward, on top of fixing the water temp itself.

Once we started doing scheduled pair retirement/replacement at 6-7 months, our average litter size per cage per month stayed flat instead of tanking.

Two things fixed it for us: 1) Switched from standard cotton nestlets to the compressed cotton nestlets PLUS a red plastic dome/hut in the cage before the female was due. Give the red dome + minimal handling a shot, it's a pretty low-effort fix.

Another postdoc in the building mentioned that C57BL/6J females can be picky about cage mates and that switching to monogamous pairing (1 male : 1 female) instead of a trio often fixes the problem, since trios can lead to social stress that suppresses receptivity. We also started doing the plug checks right at the start of the dark cycle instead of the next morning, and swapped in slightly soiled bedding from a proven breeder cage to encourage pheromone-driven cycling (Whitten effect).

Thanks everyone for the input, saved us from losing a good stud male. Taro

We resolved the C57BL/6J cannibalism issue by introducing three husbandry modifications. First, we placed two extra Nestlets (shredded cotton sheets) in the cage one week before birth, allowing the female to build a deep, enclosed dome nest. Second, we prohibited cage cleaning and all handler disturbances from day -2 of gestation through day +4 post-birth. Third, we supplemented breeder food with high-fat diet pellets and a small handful of sunflower seeds starting three days before parturition to satisfy maternal nesting and foraging behaviors. These interventions increased pup weaning rates from 45% to over 85%.

Alternatively touchdown PCR (starting a few degrees above your primers' Tm and stepping down over the first several cycles) works well too if you don't want to commit to one fixed temp, and tends to be pretty forgiving for suboptimal primer designs. Switched our SOP for this primer pair to 62C annealing and it's been consistent since.

Two changes fixed it completely for us: 1. Switched to filter (aerosol-barrier) pipette tips for all genotyping PCR setup, especially for master mix and template addition steps. Curious if it fixed the ramp rate issue people were complaining about.

We switched our protocol to an overnight digestion (12-16 hrs) at 55C using freshly aliquoted Proteinase K, and our yields went up dramatically — usually 3-5x higher than what we were getting with the 2-hour digest. Update: switched to overnight digestion with a freshly ordered batch of Proteinase K and our yields are back to normal (actually better than before).

Fix that worked for us: clean the punch tool between every single mouse — we use a quick wipe with 70% ethanol followed by a flame or a fresh disposable punch per mouse, whichever is faster for the litter size.

First, our ear-punch/tail-snip lysis buffer had been sitting at room temperature for weeks instead of being stored at 4°C, and the proteinase K in it had likely lost activity — fresh aliquots restored strong, clean bands immediately. Second, we were eluting DNA in too large a volume for old or tiny ear punches, which diluted the template below a usable concentration; cutting the elution volume roughly in half for small samples brought Ct values back into a reasonable range on downstream qPCR runs.

The thing that's worked best for us is beefing up environmental enrichment — specifically adding chew blocks and a small igloo/hut to each cage gives the non-barber mice something to redirect chewing/gnawing behavior toward, and gives subordinate mice a place to retreat to and avoid the barber mouse when they want to. We didn't need to separate the barber mouse out in most cages — just adding the chew blocks + igloo combo reduced new barbering incidents by more than half over about a month, based on our informal whisker-scoring checks.

What fixed this for us: store bags up off the floor on pallets (even simple plastic pallets work) so they're not in contact with concrete, and make sure the storage room stays below 70F — we ended up getting a small standalone AC unit for our feed closet since it wasn't on the building's main HVAC zone.

Two fixes going forward: 1. For longer transports specifically, we're switching to gel hydration packs instead of standard water bottles.

Top answer: Low humidity is almost certainly the culprit — below 30% RH, static buildup in synthetic and cotton-based nesting material makes it fray apart instead of matting into a cohesive nest, and it dries out mouse skin in the same way it dries out human skin. Two things helped us a lot: switching a portion of the nesting material to crinkle paper strips (less prone to static shredding than compressed cotton squares), and adding a small room humidifier to bring RH up into the 40-50% target range recommended by most institutional guidelines. Dandruff/dermatitis cleared up within about two weeks of the humidity fix.

The standard fix is to get room humidity up into the 40-50% range — most institutional guidelines target that band specifically for this reason, not just general animal comfort. Requesting a portable humidifier for the affected rack today while we wait on facilities to fix the HVAC long-term.

To solve the Sry primer PCR failure, we optimized our protocol by: 1) Designing multiplex primers that amplify a 200bp segment of Sry alongside a 350bp internal housekeeping control gene (like Myog or Actb) to verify lysis and PCR efficiency. 2) Reducing the tail-snip elution volume to 30 microliters to concentrate the DNA template. 3) Adding 2% DMSO to the PCR master mix to help denature the GC-rich Sry promoter regions. These modifications ensure that a missing band indicates a female pup only if the housekeeping gene is present, eliminating false-negative male identification.