Culture Journal, Species: Brachionus plicatilis

General
Species:  Brachionus plicatilis
Species description:   (assumed as L strain from Reeds)
Culture source (link if possible):  Reed Mariculture
If algae, CCMP # (Optional): 
http://ccmp.bigelow.edu/
Culture Establishment Date:  6/29/2011
Continuation Date:  10/6/2011

Vessel Details
Salinity:  1.019
Temperature:    74
pH:  8.2

Vessel description:  2x2.5g buckets
Lighting description:  none (ambient light from breeding tanks which were on 12 hour cycle)
Lighting cycle: 
Aeration description:  3-4 bps

Methodologies
Split methodology: I split the culture received between 2 2.5g buckets. 

Culture medium description: 
enough Roti Grow Plus to keep the bottom of the bucket barely visible.  2x a day.
Cell count:
 (if known)

Reference links:  

Additional Information
(No Pictures or Videos in the Section Please)
Notes: 



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So I decided I would walk back through my experience with my rotifers hoping it might help someone else, as for something so 'easy' I have struggled with it.
 
I will give short background:
I ordered my original culture on 5/18 and received on 5/20.  I had setup two 2.5g buckets that overflowed into two 5g buckets, and then into waste, where I could seive out rotifers when I choose.  The idea in the back of my head was, I could do water changes and harvest in one simple step.  Add water to 2.5g and this would maintain a low density culture while moving left over water with algae to the 5g, which would eventually wind up seived as waste.
 
I started the culture, and all was going great.  I dosed the 2.5g and the 5g at about the same rate of algea 2x a day.  The count level got quite high (as I am new to this, it is all relative), but I definatly got the...wow that is alot of rotifer reactions.
 
I routinely poured in about 1g unfiltered tank water from my breeding system (probably a bad idea) to the 2.5g daily.  So roughly a 50% water change on the 2.5g which fed the 5g below.
 
Around two weeks in, the buckets seemed to start consuming considerable amount of algea and the counts seemed to be dropping.  I did not test anything, I just resumed reading and trying to follow instructions, mostly because everyone said this was so easy.
 
Eventually, around 7/1/2011 both cultures basically crashed.  I still did not test water.  Just sat a bit frustrated.

So after this experience i decided to stop doing the 2.5g to 5g method and just try it simpler.  A local reefer suggested he had kept a culture going for over 6 months by doing nothing other than add algea, very very low bubble count (1-2 bps) and doing water changes about once a month (maybe).
 
So I ordered another round of rotifers from Reed (sigh...too much in shipping)...and gave it another go.
 
This time I setup an area outside of the breeding area away from everything where I could do just rotifers.  I wasn't sure at this point if I had contaminated my previous culture or if a parameter was astray or of the crash was caused by my breeding area being at 80-82 degrees.
 
I received this culture on 6/29/2011.  I proceeded to split this culture between two 2.5g buckets in my 'fish room' that was away from everything.  I was determined to learn as much as I can and had planned on splitting this culture into the many different methods I had read about.
 
This is my posting from http://www.mbisite.org/Forums/tm.aspx?&m=31289&mpage=1
1) Culture arrived from Reed 6/29
2) Divided culture into two 2.5 gallon buckets 1.020 water, bubbles at a rate of about 3-4/second
3) Fed with Roti grow plus until very very light green through 5ml vial, very noticably green looking through water..could still see bottom (density definately was increasing quite a bit)
4) Allowed to sit, repeating step 3 every 8-12 hours as needed to maintain
5) Day 3, started two 2L cultures, one from each bucket. Feed the same as 2.5g
6) Day 4, started sieving roughly 25% of the water volume with coffee filter.  Water went back into bucket as it left seive.  food fed to DTs.
7) Day 5, allowed all buckets to sit, witht just feeding.
8) Day 6, started new culture in glass 10g.  Fed same as others
9) Day 7, started new culture in another 2.5g from one culture.  Sieved other for DT feeding. Again using coffee filter.
10) Day 8, started seeing substantial decline in population in all cultures.  2Ls crashed..nothign noticable in them.
11) Day 9, started new culture in a two other 5gs in diff room. One from each of the origianl 2.5gs
12)  Day 10, Everything looks to be crashing. End of day (now..everything is either crashed or almost crashed).  1 5g bucket seems to ahve a density of roughly 2 per ml and two of the 2.5gs are around the same.  Everything looks pretty much empty. 
 
After the advice given on this post, I split what I had left into two buckets (it was a pretty small amount at this point).  I took half the water from each bucket and put it into new 2.5g buckets and added newly mixed (aged..about 4 days) water. So about 1g into each bucket plus 1g of new water.
 
I proceded to add my 'Ultimate' from Reeds to each bucket, in pretty large amount, about 2-ml to each bucket to help ensure that I was bringing down the ammonia levels.  The amonnia levels in the bucket I tested were at 2ish using an API test kit.
 
Bucket 1 was pushed up slightly to 5-6 bps and bucket 2 was pushed to 'boiling'.  
 
By the next morning, bucket 2 had clearly completely crashed.  There was nothing left in it at all, that I could find.  Bucket 1 seemed to be ticking along slowly.  I fed and left alone.  My feeding is generally just enought to make the bottom of the bucket disappear.
 
I rinsed and repeated this effort without harvesting.  Using roughly equal portions of 'Ultimate' and my diluted algea solution, which is about 8 ml of reeds roti grow plus in a 6oz bottle.
 
Today, 7/15/2011 the one bucket seems to have recovered enough where i felt like I could harvest some and start seeing if I could get the culture going that is 'boiling'.  I did a 50% water change on that bucket and added 4 cups of water with rotifers to it from the one 2.5g bucket that is still going.
 
 
Thats is where I am at now.  The lesson at this point seems to be....keep the ammonia down.  I plan to return to trying my 2.5g into a 5g bucket method, but I want to get these cultures going again so I don't have to make Reed any richer. =)
 
 
 

So far I seem to be doing much better.  I received my cloram-x and have been using that with 16tsp to 1L of water and dosing that at 2x the amount of paste.
 
I have done 1 50% water change and bucket change (at same time) since last post.  the roifer reproduction seems to be higher in the lower bps so far.
 
Now the trick, I have to leave again for 4 days and my 'fish sitter' had a death in the family so I don't have anyone to feed these things.  I don't have any fry younger than 2 months, so I am ok with them making due, for the few days but the rotifers will...well be dead.  So...I am going to bottle and fridge some.  In addition, to feed the cultures, I built a silly little DIY feeder with an aqualifter, timer and water cooler.  Hope it works ok.  Figured I would thin the rotifers out quite a bit, change the buckets one more time and hope for the best.  I had to leave this last week for 2 days, but I was within travel distance and I had simply overfed to start with and there was just enough algea when I got back.  Not clear, but not green either.
 
The DIY has a diluted mix of cloramx, replacement water and roti grow+.  Ice in cooler to keep it cool and the timer is set to feed 3x a day.  The valves are set to what I 'think' is a reasonable amount of food and should divide the food in the bottle by about the amount of time I will be gone.
 
Trying it out tonight, as this is my last night before I have to leave.

When I was away for an extended period of time I tried a few things. Over feeding, cold storage, etc. I find that it's a good idea to try a few things just in case something works better for you than another.

In my case, putting some in the fridge with phyto did the best. I was away for much longer than 4 days though. 

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

Well after my trip, which was rather interesting...as we lost power for about 10 hours of the time I was away...just not all at once.  From what I see the longest one was about 6 hours.  I was more worried about the fish than the rots...but I got back and all was ok.
 
So I went through the work of creating my lil DIY feeder and testing it.  It worked great during my 1 day test.  So i set it up for my trip,  Well, for whatever reason it didnt work at all.  The feeder bottle was still full.  So the cultures ran with what they had for the 4 days.  They seemed ok.  I would not say that the density had really increased at all in any of the cultures, but they survived.
 
I had added about 2x the normal feeding just before I left to get the water pretty green.  I also stuck a bottle in the fride with a pretty low density in it and added quite a bit of algea to it.
 
When I got back, did water changes on the cultures, fed again and took the bottle out and started a new culture off of it.  Three days later is looking just fine.
 
i have been doing a bucket change and 50% water change at the same time roughly every 5-7 days or so and the densitites are looking good and I am harvesting quite a bit every day.
 
So so far all is good.

The recipe for Cloram-X is to add the powder to water to make 1L, not put into 1l of water.... but it won't effect it much.
 
RG+ isn't a good choice for what you are doing.  The best choice is straight nanno.  RG+ will break down in your diluted mix.  You should never dilute then store any of our products.

Sorry, maybe wasn't too clear.  I am making a bottle of cloram-x with DI water with a mix of 16 tsp cloram-x to 1L water. 
 
I am not putting the RG+ in the cloram-x bottle.  I just did that for the few days out of town, which didn't work anyways.
 
So if I need to do some form of drop when out of town you are suggesting to use nanno instead?

We're on two different pages it appears as what you just said is what I am saying is not the protocol.
 
Your final mix should be 1 liter, not 1 liter + 16 tsp.  Take 16 tsp powder, place in 1 liter container.  Fill that container with enough RODI to make 1 Liter total of solution.
 
 
RG+ is an AMAZING product but it's not as forgiving like just straight nanno, especially in how you wanted to use it.  Your not really looking for enrichment, or any really culture expansion, just culture stability.  The great thing about Nanno is you can keep that stuff in the freezer for a long time. 
 
As for dilution and then storing of algal concentrates.... it's not a good idea.  The only time you should dilute our algal concentrates is just prior to use.
 
 
 
 

You can refrigerated rotifers in breathable bags with some nanno for a week and still have a decent culture.  I can get some numbers tomorrow on survival rates and length of time.
 
I've been telling RMI that we need to make our breathable bags available to the public.  I think they'd really help you guys when you need to do something like this.  Ours are stronger then the Kordon ones and they are free of any writing (for now) Don't get me wrong though, the Kordon ones are still pretty darn good.
 

my cultures did ok while i was gone, but I am sure they wouldn't have much longer.  I did refridge a bottle and it recovered fine after I got back.  I was looking at the breathable bags.

I've had bad experience with the Kordon breathable bags leaking.  Someone sent me a fish once in one that caused a mess for FEDEX, and a very stressed out fish.  I have stored rots in the fridge in a GLAD fresh protect bag, and they did very weill.  Still, I would not ship in one. I would be interested in seeing the ones RMI has and any info on how they make them better.
 

well these cultures seem to be going just fine.  I have been maintaining three 2.5gs now for awhile.  I have a few buckets crash, but as a result of seeing how long I could run on just cloram-x without water change.  It seems once I hit about 7-10 days I get a crash if I don't do a bucket change with about a 50% water change.  If I only do a 50% I find that the time seems to shorten before a crash.
 
Continue to harvest 1-2 a day, roughly 25% at each harvest.
 
I have contined playing with another bucket with much lower aireation and I consistently find that I am able to maintain a denser culter without it crashing with low bubble count.  I am not sure why this is exactly, but the higher the buble rate, the more likely a crash..even with densities being about half.

I am feeling more comfortable now with the cultures and my process.  I have reduced my bubble rate down to a slow bubble and this seems to be doing much better.  As I examine the cultures under scope, I only find a handful of females carrying eggs, but the cultures seem to be remaining pretty dense.
 

Pics of rots:
 

Scope pictures.
 

 

As I have been out of town alot lately, I started up a little experiement just to see what would happen while I was out of town.
 
I took 200ml of pretty dense Nannochloropsis oculata culture and placed into 600ml flask.  I filled to the 600ml mark with freshly prepared salter water matching the 1.019 salinity of the culture.  I added roughly 200 rotifers and put rubber stopper on.  Supply about 1 bubble per 5 seconds and left it alone.
 
I did this with two flask.
 
i didn't document the date as I expected them to both be totally crashed in a few weeks.  It has been at least 2 months now (started about 12/15) and the cultures are still running.  They ramped up to probably 50/ml before exhausting the visible signs of Nannochloropsis.  This took roughly two weeks.  About a week later one of the cultures appeared to have completely crashed.  Crystal clear water and no visible signs of movement.  The other flask tapered off to about 3 rotifers /ml.

Currently:

The culture that appeared to ahve crashed recovered and is now very similar to density as the other.  Both flasks have had no additions of anything.  They are holding very steady at about 3/ml.  Low density but zero mainteance.  Perfect maintenance culture.

Temp in room is pretty solid 78.  Flasks sit next to the rest of the phyto cultures in front of dual T5 lights.