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Post by lenynero on Oct 27, 2009 14:16:46 GMT -5
Actually Hackerberry gave me the link ;-)
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Post by lloyd on Nov 2, 2009 21:26:18 GMT -5
My HEPA filter is 16" x 20". My feeling is that the working area should be about the same depth as the width of the filter or 16" in my case. Also I think the part in back of the filter should be enough so that the fan (an inline centrifugal fan) will comfortably sit inside. Any thoughts? Also my filter has a little band of dead space around the periphery where the filter part is cemented to the case. Do I have to exactly match the working area to the active area or just have it be the same dimensions as the filter case? I would imagine that having it exactly the same as the active part of the filter would be better for good laminar flow but maybe it doesn't matter?
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Post by tom on Nov 15, 2009 11:18:00 GMT -5
Sorry for the late answer Lloyd, but here i am I'm not sure to underdstand all the subtilities of your questions but despite im not a pro in LFH dynamics, i think that: -your working area doesnt need to be as deep as the width of the filter. A good example of this are the 'dual work place' LFH used in lab, where the stainless steel work place is about 4' wide but about 2' from your chest to the filter. You have to work as safely close to the filter as you can (when possible, depending of your 'sterilizing agent (open flame, bacticinerator), if you can't quantify where your sterile area stop with your wind velocity/and if its a turbulent/laminor air flow. -about the room for the fan: i agree that the fan should fit confortably in its dedicated space. Should be easier for you to move/tickle with it if needed (repair, etc) and might help about cooling some parts? I dont think there is some rectriction about the size of the 'room' it should sit in, the 'in' and 'out' hole dimension should be what will drive the fan capacity/ease to create a proper airflow. -All filter that i have seen have the 'deadspace' around the filter (about 1"). You dont have to fit it perfectly with the working area. Since you'll be most likely spraying the area with rubbing alcohol and make the LFH run for 10-15 min before working under it (it 'cleans' the air of the working area, and the alcohol would do the job for the working surface and air deadspaces). i wouldnt worry about it. Once again, the ones i've worked with in labs had this 'dad space' that i could see. If this still worry you, but its too much trouble to fit it perfectly, assuming that your airflow is laminar, just just work at the height of the dead space, which would be impratical anyway since they are located at the 'border' of your sterile area. -the airflow is, by what i recall, more about how the air will act once getting through the filter and might be more about its velocity (and influence from air current in the room, if there is strong ones) than the dead space around the filter. Of course, if your working space is 'enclosed' (i think Renesis' one was like this), ie a bit if you are working in a corridor like area in front of your filter (vs a filter 'alone), it might help to preserve the 'integrity' of the air flow. An easy way to tell about would be a flame test/smoke test, ie run your LFH and place a burner or a candle in it and try it at various place while observing the flame or smoke behavior. If it lays on its side without moving too much, i think it would be suitable. It can help to predict as well at what distance of the filter you should work within to be in the 'good air flow' area. In the same 'state of mind' in the case of a too powerful fan vs filter size, a rheostat would be interesting, so you'll be able to gauge the fan speed for a proper flow and with a maximum or working area. I think there was a documents about the airflow/movement in the docs i sent you a while ago, but im not sure about this. I might try to find it back on the web. Sorry for all the '' ''... i hope the message can be understood Tom, lacking his coffee update: some links www.globalrph.com/aseptic.htmairforcemedicine.afms.mil/idc/groups/public/documents/afms/ctb_026442.pdfhope this help
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Post by lloyd on Nov 15, 2009 23:11:36 GMT -5
Once again, Professor Tom carefully guides his TC acolytes.....
Thanks, Tom.
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Post by lloyd on Apr 7, 2010 13:31:18 GMT -5
I finally finished testing my home made Laminar Flow Hood. No growth at 2 weeks so I think it works. Click on my website for photo's and documentation (if you like looking at big boxes ;-) )
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Post by hackerberry on Apr 7, 2010 13:44:04 GMT -5
Awesome!
hb
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Post by renesis on Apr 7, 2010 19:53:15 GMT -5
Looks amazing Lloyd! Great work!
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Post by lloyd on Jun 17, 2015 22:02:03 GMT -5
Still doing TC. Last few years doing orchid seeds which is fun. Now I've got the ceph seeds from Snapperhead to try. Just mixed up some Phytotech medium for them. For some strange reason I always added the agar to the medium before dividing it up into the bottles which always wastes some agar and distributes it unevenly. I looked at a web site today where it said to put the agar in each bottle which was so much easier. Live and learn.
I've also got to replate a bottle of orchid seedlings as they are going crazy, must be hundreds of the little guys.
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Post by hebtwo on Aug 14, 2015 7:28:26 GMT -5
Still doing TC. Last few years doing orchid seeds which is fun. Now I've got the ceph seeds from Snapperhead to try. Just mixed up some Phytotech medium for them. For some strange reason I always added the agar to the medium before dividing it up into the bottles which always wastes some agar and distributes it unevenly. I looked at a web site today where it said to put the agar in each bottle which was so much easier. Live and learn. I've also got to replate a bottle of orchid seedlings as they are going crazy, must be hundreds of the little guys. I'm curious Lloyd, what kind of orchids are you culturing?
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Post by lloyd on Aug 17, 2015 20:38:51 GMT -5
Luisia zollingeri
Dendrobium sutepense
Coelogyne xyrekes
Cirrhopetalum lepidum
These are the latest selections from Thailand.
I've had a lot of contamination problems lately so not a lot of success. Some bottles are growing Ok.
I think I tracked the problem down to the pressure cooker. I've been using tap water and there was a lot of mineral build up in the pressure valve. I suspect the temperatures were not high enough to kill spores. I replaced all the replaceable parts including the pressure regulator. I just did a batch today, so I'm hopeful it will be Ok.
Morale of the story: use distilled water when using the pressure cooker. I checked the water afterward and it went from 5L to 3.5 liters and from 0 to 3 PPM so you don't even use up that much distilled water. It's worth it to prevent wear and tear on the pressure cooker.
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Post by deanm on Aug 17, 2015 20:45:09 GMT -5
Although I have used a pressure cooker for sterilizing I much prefer an autoclave. 15 years ago I ran a commercial plant tissue culture lab in the lower mainland of BC (Brookside Greenhouses Ltd).
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Post by deanm on Aug 17, 2015 20:46:21 GMT -5
Lloyd is right about using distilled water - saves a lot of wear and tear for autoclaves also.
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Post by lloyd on Aug 17, 2015 21:04:53 GMT -5
The sterilizing is the easiest part, usually. It's the sterile technique that usually trips me up.
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Post by deanm on Aug 17, 2015 21:30:24 GMT -5
Yes, that is the tricky part. I can empathize with you on that.
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Post by snapperhead51 on Aug 17, 2015 22:48:49 GMT -5
what part of sterile technique is your biggest problem LLoyd ? mite be able to help out there , if if want some help
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