I'm a microbiologist recently who recently started working on an ongoing C. difficile project for the first time. One of the things I need to do is verify the lower detection limit of my qPCR detection assay (C. diff Toxin B gene) using C. difficile spores spiked into negative clinical specimens. I also need to obtain accurate viable spore CFU counts in order to generate the appropriate spore titers for animal experiments. I'm trying to get a clean spore prep free of vegetative cells (for the spore detection assay), and minimize the number of spores clumping together to obtain more accurate CFU counts (via plating heat-treated spore dilutions on BHIS+taurocholate germination plates).

I'm working with the C. difficile strain 630 (ATCC BAA-1382), a lab strain that's notoriously not the best sporulator (we use 630 because it's toxigenic and infectious, but not highly virulent. This helps minimize animal suffering in simple infection models where we don't need to look at toxicity).

There are a few commonly cited C. difficile spore prep methods that I've been working from, and in some cases trying to mix and match possible solutions from different protocols, but none of them were use this particular strain in the methods papers. Our lab SOP uses Clospore medium, and that seems to give me the best sporulation rates compared to the solid 70/30 medium used in the STAR protocols paper. But when I follow the rest of the Clospore protocol for spore collection and purification I end up with highly concentrated vegetative cells and am getting lower viable spore titers in the "concentrated" spore prep than there are in the unconcentrated cultures (by multiple orders of magnitude). The spores I do recover are mostly clumped together into large clusters when viewed under phase contrast microscopy (possibly explaining the lower spore CFU titers).

If anyone has experience with getting really clean C. difficile spore preps with minimal spore clumping (especially in 630), I would love any tips or advice.

Potential solutions that I've tried without success:

  1. Saving the pellet wash supernatants and re-pelleting with a longer centrifugation: This does recover some spores with almost no vegetative cells, but they are very low titers and still have clumping issues.
  2. Using room temperature water instead of cold water for washing the spore pellet, (it's been reported that cold-water wash induces more clumping): Mine still clump with room temperature water, and there's no difference in terms of vegetative cells.
  3. Separation of spores from vegetative cells using a 50% sucrose gradient centrifugation: This does separate out a lot of cellular debris, but the intact vegetative cells pellet out with the spores, and still has spore clumping issues. Plus I'm losing a lot of spores when I do this (possibly due to extra wash steps needed).
  4. Washing with 0.05% TWEEN/Polysorbate (nonionic surfactant): This actually helps to free up spores sticking to the culture tubes, but they still clump together when washed to remove the TWEEN (polysorbate has toxic effects in mice, so washing it away in not optional).
  • $\begingroup$ I’ve had similar poor luck getting sufficient spore concentrations following this protocol and using ATCC strain 9689. Lots of vegetative cells, few spores. One recommendation from that protocol is to use PBS + 1% (w/v) BSA to resuspend spores after washing to prevent clumping, per Edwards et al. $\endgroup$
    – acvill
    Jul 4 at 14:24
  • $\begingroup$ @acvill Thanks! I'll will try that for sure. $\endgroup$
    – MikeyC
    Jul 6 at 14:47

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