Live Algae vs Algae concentrates

[clownfish75]

Hi All

I was just wondering who has used both live and dead algaes in their larval rearing tanks (green water techinique) and wondering if they found any difference in survival.

I origionally started with live nanno 4 years ago but moved to the reed concentrate nanno/roti diet in the last few years, due to price im trying ot get back to live.

the questions i have are, that my dead algae systems have a .5 pH movement over a day would the live stuff stop this, and has anyone ever compared the 2 in that respect.

Im not concerned with ease of use etc, more specifically the function properies.

Christian

 

[Dman]

That's funny, our production meeting today centered around that very topic (time to re-stock the IA reserves) and I'm about to embark upon an non-scientific study on various algaes in co-culture to determine which is a better route. The outcome will be centered around cost vs. survivability to post-meta.

I'll be back in a couple of hours with a brief outline. Have to get back to work.

 

[mwp]

FWIW, live algae has one distinct advantage over "statis" types...live algae can take up waste nutrients. O2 produced during the day too, but at night CO2 produced, which could cause pH swings unless aeration is vigorous. I've always been worried about pastes and such "spoiling" or otherwise breaking down in the tank when it comes to using them for Greenwater Techniques, although more than one source has suggested that isn't a problem...

HMM.

I like live, it's relatively cheap to produce and in the case of MOTILE species (T-Iso, Tetraselmis) it may even be a direct "First food" for the smallest of fish larvae.

I'm VERY interested to hear what Dman comes up with.

FWIW,

Matt

 

[GreshamH]

I'd think that live will have a greater swing. We have to beat down our phyto PH with major automation. A phyto culture can get far to high for most fish if it's booming. Of course I'm biased though IA has got to be on the pricy side in Australia.

We've got many studies on this very subject from major fish farms from around the world. I'll see if any aren't copyrighted so I can post them here

 

[clownfish75]

Thanks guys, great to hear info on exactly what i wanted, IA is about $150 a litre local currency, currecny exchange is about 3/4 that in US dollars.

With the IA i get a .5 swing down during the day, at nightt he system is filtered so the live algae would not get a chance to produce CO2 so much ar night. And certianly not to the detriment of the larval tanks.

The other thing i have been considering is the rotifer density i know someone who did a small experiment and found that anything from 10 rots per ml all the way upto 100per ml got great survival with increasing density, so maybe at higher densities nitrogen production is up so maybe live algae would have some sort of filtering effect.

Ill take some samples over the day soon to see if it degrades with high rotifers and IA.

Christian

 

[Kathy55g]

Great idea. I am particularly intereseted in the increased concentrations of rotifers. My rotifer densities tend to disappear soon after I add them. This is particularly true with large batches, so I am thinking that they are just getting eaten. Sometimes, though, they sink to the bottom of the tank and do not swim as much as I would like. Perhaps some change in salinity issue there. Would adding more be harmful, or beneficial?

 

[ediaz]

I don't see any difference with any specie i work with, besides that harvesting,bleaching, sterilizing and coming from work to wonderful crashes versus giving delightful Kathy a call, and open the fridge to green fresh ultra high density algae.

I use about $70 a month of it thats $17 a week, my time is more valuable than that.

I don't know of any species that are being produced to date that require phyto mixes or motile algaes unleess someone is working on something I don't know.

Ed

 

[mwp]

Mandarins baby!

Matt

 

[ediaz]

"being produced "

my ex girlfriend says hi

 

[Dman]

I'll have to keep this short and sweet, as it's almost time to get back to work.

I plan to loosely quantify the following parameters with different algae’s; live, dead or otherwise and a combination thereof using my WSM hatches as the test subjects.

In each module I will log the following:
Approximate clutch size
Percentage hatch
Percentage caught

Daily water parameter logging:
Temperature
pH
Salinity

Qualitative daily rotifer densities:
Light, medium, dense
And any rotifer additions
(Sorry, I'm cute not technically savvy)

Qualitative water colour as it pertains to algal concentration:
light, medium, dense
(Once again, see explanation above)

Qualitative measurements of aeration:
Light, medium, heavy.

Daily 2 gallon water changes with bottom siphons will be used to count the expired larvae and replacement water will consist of 1 (one) gallon parent water and 1 (one) gallon ASW mixed and aerated a minimum of 24 hrs. and adjusted to 1.018 salinity and 8.00 pH. (Here's a question I meant to start as another thread, should I drip the replacement water in over a period of an hour or pour it in in such a fashion as to not damage the larvae?)
Also included in the replacement water will be 0.2ml of Lugol's at 5% w/w and 0.1mg CloramX

Algae additions will be:
1. Live Nannochloropsis Occulata
2. Dead Nannochloropsis Occulata (IA)
3. Live Nannochloropsis Occulata and live Isochrysis
4. Dead Nannochloropsis Occulata and live Isochrysis
5. Live Nannochloropsis Occulata and live Tetraselmis
6. Dead Nannochloropsis Occulata and live Tetraselmis

I'm limited to these three live algae’s as they're the only ones available to me locally.
(Again, I’m cute not savvy so I’m not counting freakin’ cells)
The idea is to see which of the above combinations have the best overall survival rates. Any oversights, f-ups or catastrophic deaths during a test module by either myself, my partner or Mother Nature will be repeated to remove said screw-ups from the test results.

Equipment

10-gallon tanks are all glass and are uniform in size and will be filled with 5 gallons parent water filtered through a 53 micro screen the morning following the hatching event.

Aeration will be supplied to each of the test tanks by a Hagen Optima air pump and regulated with the use of brass manifolds.

A Hanna hand-held pH and temperature meter will be used to test, ummm pH and temperature.

A hand-held refractometer with ATC will be calibrated daily and used to measure salinity.

Is there anything I should add? (Keeping in mind I have a wife, two kids, a day job and I’m also not looking to put together a Master’s thesis here)

 

[mwp]

I think rotifer densities and additions should be measured for this to have any real meaning, sorry, but that's my $0.02. Ideally of course you'd want to count EVERYTHING and quantify everything, but rotifer densities are consistently cited in literature etc, so there's a basis for suggesting that they are important enough to measure.

Algae densities would be KINDA easy...not saying you have to actually count them too, but you could use a Sechi Disk (spelling) to quickly get a basic "density"...granted a measurement in a tank getting only Nan would't be the same as a nan-t-iso mix, but it WOULD show relative increases and decreases within each over time. Granted, if you know what ratio the algae are being mixed in, you could probably come up with actual density estimates for the mixed feeds...

I just think if you're going this far, it may not hurt to go just a bit further and get the hard data...makes it more difficult to discount or dismiss the findings.

FWIW,

Matt

 

[Dman]

Quickly,
If anyone has the means or the clearance, check to see if this has been done?
Or something similar?
Or perhaps with a different species, I'm sure someone's probably done this with finfish.

 

[Dman]

Quickly,
If anyone has the means or the clearance, check to see if this has been done?
Or something similar?
Or perhaps with a different species, I'm sure someone's probably done this with finfish.
As I have no idea where to look

 

[mwp]

BTW Dman, you can get ACTUAL clutch size if you can take pix of the eggs right before they hatch. A couple of us might be interested if you took pics of the nests when LAID and then right before hatch, to see what percentage of egg loss you have during incubation, but that's a whole 2nd study And besides, if I know anything about maroons, I'd be thinking that getting a picture of the entire egg clutch in focus and large enough for counting, might be problematic in itself. But you did ask for suggestions / opinions, right?

Matt

 

[GreshamH]

<a href=showthread.php?s=&postid=9026809#post9026809 target=_blank>Originally posted</a> by Dman
Quickly,
If anyone has the means or the clearance, check to see if this has been done?
Or something similar?
Or perhaps with a different species, I'm sure someone's probably done this with finfish.
As I have no idea where to look

We've got many studies on this very subject from major fish farms from around the world. I'll see if any aren't copyrighted so I can post them here

Every major hatchery that switches to IA paste does such studies. They tend to be for in house research though and we all know how loose lipped hatcheries are with their research :lol: I do recall one in a EU aquaqculture mag though.

I've been out off town all week, so I have not had a chance to do what I quoted above Monday for sure

 

[GreshamH]

This is sort of on subject, if you can get the paper.

DEVELOPMENT OF THE PH IN THE INTESTINAL TRACT OF LARVAL TURBOT
K. Hoehne-Reitan, E. Kjørsvik, K.I. Reitan-2001
Marine Biology, 139(6): 1159-1164
Abstract:
The pH in the gut of turbot larvae and juveniles of turbot was studied from day 11 until the completion of metamorphosis. Dietary effects on the gut pH were estimated when larvae were offered live feed, a microdiet, only microalgae or no feed. The pH in the gut was weakly alkaline until day 24 after hatching with no differences between the foregut, midgut and hindgut. The foregut contents started to turn acidic from day 28 after hatching when the larvae were already weaned successfully, which indicates that an acidic pH is not necessary for the digestion and utilisation of formulated feed. During the following 20 days the pH in the foregut/stomach decreased further to a minimum of pH 3.5, while the pH in the midgut and the hindgut increased slightly to a maximum of pH 9.0. Larvae receiving live feed, microdiet or microalgae had a similar pH in the midgut on day 11, while starved larvae exhibited a lower gut pH. This suggests bicarbonate secretion from the larval pancreas stimulated by ingested microalgae or feed particles.
(Norwegian University of Science and Technology (NTNU), Department of Zoology, Brattøra Research Centre, 7491 Trondheim, Norway, Tel: +47-73-590321, Fax: +47-73-596311, E-mail of K.I. Reitan: Katja.Reitan@chembio.ntnu.no)

 

[Dman]

Gresh, thanks.
Quick update, had a spawn last night and a ten gallon tank will not be big enough. So I'm upgrading tank size to 15 gallon tanks with ten gallons of parent water.

I'll also get a more accurate count of rotifer densities by physically removing and visually counting one (1) millilitre of tank water at midday. (Which would be 8pm)

When I have a couple of days worth of data I'll start a log a la MWP.
Any suggestions regarding reporting format. I'll be recording everything here locally on an Excel spreadsheet.

 

[Dman]

Quick update.
I started with a WSM hatch that numbered approximately 2000 eggs. Of which I captured 80% approximately.
Honestly, I think I'm going to have to pick a more appropriate pair to work with, or at least an easier pair. This is day two and there's no way I'm going to be able to siphon their tank today as there is literally a cloud of fry in the tank. I abandoned the bottom siphon as there were almost four times as many alive scurrying along the bottom as there were dead ones. Of the 100ml I siphoned there were 15 dead and almost 57 that I had to pipette out and return. (this took almost an hour)

Initial water parameters:

01/22/06 22:00 EST

pH 8.03
22.1 C
1.018 salinity

Rotifers:
10/ml

IA addition:
1 ml or +/- 68 billion cells

Water parameters:

01/23/06 20:00 EST

7.90 pH
21.8 C
1.018 salinity

Rotifers:
8/ml

Additions:

Rotifers:

 

[Dman]

<a href=showthread.php?s=&postid=9058897#post9058897 target=_blank>Originally posted</a> by Dman
Quick update.
I started with a WSM hatch that numbered approximately 2000 eggs. Of which I captured 80% approximately.
Honestly, I think I'm going to have to pick a more appropriate pair to work with, or at least an easier pair. This is day two and there's no way I'm going to be able to siphon their tank today as there is literally a cloud of fry in the tank. I abandoned the bottom siphon as there were almost four times as many alive scurrying along the bottom as there were dead ones. Of the 100ml I siphoned there were 15 dead and almost 57 that I had to pipette out and return. (this took almost an hour)

Initial water parameters:

01/22/06 22:00 EST

pH 8.03
22.1 C
1.018 salinity

Rotifers:
10/ml

IA addition:
1 ml or +/- 68 billion cells

Water parameters:

01/23/06 20:00 EST

7.90 pH
21.8 C
1.018 salinity

Rotifers:
8/ml

Additions:

Rotifers:

This is what got missed as my 'puter decided to crap out on me

Rotifers:
8/ml

Additions:

Rotifers:
7/ml to bring up to 15 per milliliter.

0.5g CloramX

6ml Randy’s Baked Baking soda solution

The 100 ml siphoned out was not replaced. Rotifer densities are an average of five 1ml samples taken from various area of the tank and visually counted.

 

[Dman]

Unit 1, Day 3

01/24/07 20:00EST

Measurements and Observations

7.80 pH
26.6 C *
1.018 salinity
Rotifers: 10/ml
Water was pretty much almost clear tint-wise

Water siphon

2 gallons out and subsequently replaced with 3 gallons parent water. I initially started out bottom siphoning and again this turned into an exercise in futility. I had siphoned less than 2 cups of water and had acquired 17 dead and 42 alive that needed to be returned. (Another hours’ work) The remaining water was siphoned through a 53-micron sieve to keep rotifers and larvae in the tank.
The 3 gallons of parent water was added as we determined that there was too many larvae even for 10 gallons and that we would over a period of a couple days increase the volume in the tank.


Additions:

Rotifers:
5/ml to bring density up to 15 per milliliter

0.5grams CloramX

½ cup Randy’s Baked Baking soda solution

1ml IA Nanno 3600

* A heater was added the previous day and the temperature was allowed to drift up over the course of 18 hours