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Rainwater from a butt - de-oxygenated?

Hi all,
The rainwater from the butt measured only 2 mg/l, which is the lowest level that the kit can indicate. For water at outside temperature (probably about 10°C), that is very low. Given that this is the lowest level the kit can indicate, it's possible that the rainwater is even lower in oxygen than this.
Does the butt have a sealed top? The water in the butt should remain oxygenated, if there is access to atmospheric gases, assuming that there isn't a lot of organic matter in the bottom of the butt (which would decompose).

Even though I have diverters, or a strainer, on the down-pipes, I still tend to get bit in the water butts (mainly moss and lichen from the roof) and I give them a clean out every 18 months or so.

cheers Darrel
 
Hi Darrel - yes, the butt does have a lid. Perhaps I'll try leaving it off and see if that makes a difference.
 
Ottos can breathe from the surface and if oxygen is the issue, they'll be the least stressed ones in the tank, plus they'll shoot to the surface to gulp air.
I'm confused on this ... it's my understanding that oto's gasp surface air to fill "sacs" that are then used to control "flotation" (I'm saying this all rather badly) but they are unable to actually utilize that stored air for oxygen.

Aplogies Dr Mike Oxgreen for the hijack :sorry:
- rather more on topic, do you add the water with splashing? also filter surface movement?
 
I think some fish can gulp air into their digestive tract and the blood vessels can take some oxygen from that air; I don't know if loricariids such as Otos can do this.

When adding water back during a water change I tend to pour the water over my hand to minimise disturbance to the tank. My filter return is via a spray bar mounted low down, pointing upwards up the rear glass. This gives very gentle flow and agitation, which I think is in keeping with the chili rasboras that I'm keeping. Provided I keep my stem plant (Pogostemon erectus) pinched back so that it doesn't block the water surface, I tend to get enough forward flow across the surface to transport CO₂ to the front and somewhat downward to my carpet plants.
 
I'm confused on this ... it's my understanding that oto's gasp surface air to fill "sacs" that are then used to control "flotation" (I'm saying this all rather badly) but they are unable to actually utilize that stored air for oxygen.

Aplogies Dr Mike Oxgreen for the hijack :sorry:
- rather more on topic, do you add the water with splashing? also filter surface movement?

My apologies for your confusion.
I am not entirely sure of the biological mechanism but they can utilise aerial oxygen. In nature they are found in rather stagnant conditions as well. Here is a random scientific paper about it:

"Loricariid catfishes have evolved several modifications of the digestive tract that


appear to fWIction as accessory respiratory organs or hydrostatic organs. Adaptations


include an enlarged stomach in Pterygoplichthys, Liposan:us, Glyptoperichthys,


Hemiancistrus annectens, Hemiancistrus maracaiboensis, HyposWmus panamensis, and

Lithoxus; a U-shaped diverticulum in Rhinelepis, Pseudorinelepis, Pogonopoma, and Pogonopomoides;

and a ringlike diverticulum in Otocinclus. Scoloplacids, closely related

to loricariids, have enlarged, clear, air-filled stomachs similar to that of Lithoxus. The


ability to breathe air in Otocinclus was confirmed; the ability of Lithoxus and Scoloplax

to breathe air is inferred from morphology. The diverticula of Pogonopomoides and

Pogonopoma are similar to swim bladders and may be used as hydrostatic organs."

source: http://www.auburn.edu/~armbrjw/Air.pdf
 
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Thanks for the link ... still don't see how they actually proved anything though ;) - reasoned supposition based upon morphology isn't quite the same

Some of my scepticism is watching my own oto's - after they rise & gulp air, then return to lower in the tank, I still observe expected gill movement so I'd want more evidence than the author's contention that he did not observe gill movement in his own oto's after surface gulping :)
- note that I seldom observe the gulping behaviour.

I did recently observe a severely distressed oto at the surface on it's back gulping air, then rolling, returning to inverted position, more gulping ... all the while also pumping gills - this poor fellow arrived in a recent shipment that did not go well (bags fouled etc though shipping had proceeding according to schedule) He was DOA an hour or so later.
Very few of the otos in the shipment engaged in the surface behaviour - those that did were among the most stressed (& mostly dead :sorry:)



Because of the different shapes of the diverticula


and the notion that Otocinclus and the

Rhinelepis group are not considered to be closely

related (Isbriicker, 1980; Schaefer, 1986, 1990),

state 7 is not considered to be homologous to

states 4-6. That Otocinclus nearly stop irrigating

their gills when they commence gulping air supports

the contention that they are air breathers.

However, it does not prove that Otocinclus are

using the diverticulum to extract oxygen; they

may be using the diverticulum only for hydrostatic

control. The diverticulum in Otocinclus has

extremely thin walls (much thinner than airbreathing

structures in all other loricariids), further

suggesting the use of the diverticulum as a

hydrostatic organ.
 
Hi all,
The ability to breathe air in Otocinclus was confirmed
Thanks for the link ... still don't see how they actually proved anything though
I've got the reference from the Armbruster paper somewhere, it is from an un-published Ph. D thesis.

Jay Nelson at Towson has done some work on <"quantifying air breathing in Loricariids">, but I don't know if he has worked with Otocinclus (and I don't have access to the paper).

The ability to extract oxygen from atmospheric air is thought to be derived from the common ancestor of the Loricariidae and Callichthyidae (they are both families in the superfamily Loricarioidea), but it has been lost in rheophilic species (Hypancistrus spp. etc). They still gulp air in times of oxygen stress, but they can't extract any oxygen from it.
- note that I seldom observe the gulping behaviour.
I haven't either, but I know from threads, on PlanetCatfish etc., that people who keep Otocinclus in less well oxygenated tanks assume that it is standard behaviour.

cheers Darrel
 
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Thanks for chipping in Darrel :D

(hmmm not sure I said that right :oops: - contributing, I mean)
 
I reckon it could well depend on the species...mine have always darted up to the surface to gulp air even in my low-energy tanks which have always been very well oxygenated.
Like SF has said apparently the digestive tract adaptations that allow them to "air breath" is a diagnostic trait of the genus.
Loricariids are mostly facultative air breathers but I've heard/read somewhere that may not be true of all oto species...in that some maybe obligate air breathers
 
Otocinclus spp. A duct forms at the junction between the esophagus and the stomach and expands into an enlarged, ring-like diverticulum, diagnostic of this genus, which allows air-breathing.

Personally, during the winter I may not see otocinclus gulping air at all. In the summer when my tank's temps go up to 28-29 they start doing it and it is then I start worrying because it's a sure sign of low oxygen. As Troi says they are probably facultative breathers. They do survive in stagnant waters in dry seasons alongside corydoras species so they must have a mechanism to extract oxygen.

Here in this video Ivan Mikolji mentions otocinclus and corydoras in relation to their very high tolerance of low oxygen conditions. (at around 11:20) He's filming pools of water created by the preceding rain season which eventually get cut out from the main river. He says that normally the only species that remain are those tolerating these very low oxygen water pools, mentioning the air gulping behaviour of corydoras and otocinclus.




And here is another one of locals catching otocinclus in those same stagnant pools before they dry out completely. Look at the amount of otos in one catch...!

 
When doing my 50% water change, I use 9 litres of rainwater from my water butt, mixed with 3 litres of unsoftened tap water. This gives me roughly the 3° KH and 6° GH that I'm after, and I can tweak the GH up a bit by adding MgSO₄ solution if necessary. I use Seachem Prime as my dechlorinator, and use a full-tank dose (only 0.5ml for my ~20 litre tank, so it would be impossible to measure any less). I boil a litre or so of the rainwater to bring the new water up to temperature.

I often notice that the gill rate of my Otocinclus has risen dramatically after a water change, and this seemed particularly bad yesterday. They were also very restless, and the chili rasboras were nervous as well - schooling very tightly and patrolling the tank rapidly. Most of the blue cherry shrimps seemed fine, but I did see one swimming very erratically and seeming to have difficulty staying upright. There also seemed to be a migration of snails heading up the glass towards the surface, within half an hour of the water change.

I decided that there might be a problem with lack of oxygen in the new water, although I must admit that none of the fish were gasping at the surface. To combat this, I added 15ml of 3% Hydrogen peroxide in the hope that it would provide some emergency oxygenation. (Actually it may have been after this that I saw the shrimp swimming erratically, not sure). After this, the chilies seemed to calm down slightly over the course of the next hour or so, and perhaps the Otos' gill rate slowed very slightly although still too fast.

So, does anyone know the typical oxygen content of rainwater that has been sitting in a water butt for a few weeks? I can't decide whether it's likely to be de-oxygenated or not. I am drawing water from the bottom of the butt.

Is there any point trying an oxygen test kit?

I can't think of any other contamination: the water butt is plastic, as is the house guttering. The roof tiles are old, so shouldn't be leaching anything into the water any more. The bucket and jug that I use are not used for anything else. That said, I don't know if it's my imagination but the tank didn't look quite as sparkly clear just after the water change - but that could be normal.

The tank looks okay this morning although I haven't yet done a head count.

Any thoughts? Would low O₂ level cause rapid gill movement in one species, but 'stressed' behaviour in another species, without any gasping at the surface?

Hi Dr. + all ..

I`m new to this forum but not new to keeping tanks..
I`ve kept tanks for more than 30 years..

Have you checked the NH4+(ammonium), NH3 (ammonia) and pH in the tank + the butt..?
When changing that large amount of water at once.. the pH in the added water+tank water plays a significant role if the "resting"/existing amounts of NH4+ is high i your tank.. (A lot of "restproducts" in the tank-system so to speak..)

When/If pH raises to over 7 in your tank, the NH4+ may almost immediately turn into even worse poisonus and aggressive NH3..
..which may then destroy the gills on fishes (shrimps??) very fast, and as I`ve understood it..it is almost irreversible, .. and therefore making it impossible for most fishes to absorb oxygene from the water..
This may explain the sudden "panic" behaviours from your fishes + shrimps!!??

This same situation may happen if you get some newly purchased fishes in a very "contaminated" transportation-fish/plastic-bag..
..and are adding water with a high pH in it... acute suffication may occur !!!

This may also explain why all the Otocinclus in the film never gets bad.. even if the pool/pond is heavily contaminated(from what it looks..) ..the pH is probably lower than 7...!!!??

I don`t know if this is accurate for what happened to your tank, but it may be worth thinking of..??

/Micke
 
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Have you checked the NH4+(ammonium), NH3 (ammonia) and pH in the tank + the butt..?
NH₃/NH₄⁺ in the butt measures zero. The tank also always measures zero. I haven't measured the butt pH.

When/If pH raises to over 7 in your tank, the NH4+ may almost immediately turn into even worse poisonus and aggressive NH3..
That shouldn't be happening because I'm using Prime as my water conditioner, and dosing a full-tank dose with each water change (largely because it's impossible to measure a smaller quantity). Prime converts ammonia, nitrite and nitrate to much less harmful forms.

I never see any positive ammonia reading, either in the tank water or the new water. I rather doubt that there's enough NH₃/NH₄⁺ around to cause damage, regardless of the pH.
 
Right, I have done the water change.

I prepared the new water yesterday afternoon, using 8 litres of rainwater, 3 litres of unsoftened tap water, and 10ml of the MgSO₄ solution that I mentioned earlier (which is enough to raise the whole tank's GH by about 0.5°). I aerated with an air stone, which has been running for about 18 hours. Before adding this water to the tank, I boiled another litre of rainwater and mixed it in to bring the temperature up.

I measured the O₂ levels of the tank (before the change) and the new water. The tank, which has been in darkness overnight, measured only 2mg/l, whereas the new water measured about 5mg/l. I'm not sure I believe the absolute values, because surely at that level the fish in the tank would be showing signs of distress, but they were all behaving quite normally and the Otos' gill rate was slow (1 or 2 per second). However, I'm quite willing to believe the relative readings, namely that the new water was better oxygenated than the tank, having been aggressively aerated for a long period. I suspect this kit under-reads.

The parameters of the tank water before the change were: Temp 25.0°C, TDS 139, pH 6.5, NO₂⁻ 0, NO₃⁻ 40, KH 3°, GH 5°, O₂ 2mg/l

The parameters of the tank water after the change were: Temp 24.7°C, TDS 121, pH 7.0, NO₂⁻ 0, NO₃⁻ ~30, KH 3°, GH 6°

So, there is a modest reduction in TDS, accounted for largely by the reduction in nitrate (and probably other excess nutrients as well). This is, after all, the whole point of changing water - a moderate reduction in TDS seems inevitable when doing a water change.

The pH change is inevitable, since I'm adding CO₂ depleted water (the drop checker always goes a bit bluer after a change). Probably the removal of organic wastes would also contribute to this - which is, again, the whole point of changing water. I don't see that there's much that can be done about that.

I didn't bother re-measuring the oxygen content because I know that it would have gone up, although only slightly and I don't think the test kit would be able to indicate the change.

It's too early to assess the behaviour of the chilies, because the lights are still off - but within minutes of refilling the tank, the Otos' gill rate had risen sharply, although perhaps not as dramatically as last week and they're not behaving unsettled.

I'm beginning to think that this is just a slight 'panic' reaction in the Otos. It's certainly true that they do sometimes panic and swim about wildly when I'm pruning plants etc. Last week it may have been worse because I did the water change with the lights on.
 
Hi all,
I measured the O₂ levels of the tank (before the change) and the new water. The tank, which has been in darkness overnight, measured only 2mg/l, whereas the new water measured about 5mg/l. I'm not sure I believe the absolute values, because surely at that level the fish in the tank would be showing signs of distress, but they were all behaving quite normally and the Otos' gill rate was slow (1 or 2 per second). However, I'm quite willing to believe the relative readings, namely that the new water was better oxygenated than the tank, having been aggressively aerated for a long period. I suspect this kit under-reads.
They won't be accurate values, measuring most dissolved gases is pretty difficult. You really need a DO meter.

I would assume that your aerated butt water was pretty fully saturated with oxygen, I would be a little bit worried that the level for the tank at night was 1/2 that level. Levels in mg l-1 (ppm) dissolved oxygen can be converted to percentages if you know the water temperature, atmospheric pressure and conductivity of the water, although in fresh water you can ignore conductivity, and to a large extent, atmospheric pressure. Chart below.

nomogram.gif


We use these at the moment <"Hach HQ40d portable..">, with <IntelliCAL™ LDO101 Rugged probes>, these are the best DO meters we've had, but it is quite an expensive option. You still need to calibrate the meter in water vapour saturated air before use, but it doesn't constantly need re-calibration.

cheers Darrel
 
Actually, remembering back to the late 80's when my friend had marine fish and using rain water was all the rage, many people had issues with rain water killing fish. This was traced to leaching preservative from roofing felt/underlay, fitted under tiles in modern houses. Normally this felt/underlay never comes into contact with water, but at the lower ends of the roof, it often dips into the gutter and it was here it was leaching into the rain water. The collected water was fine for plant watering but fatal to fish.
 
I would assume that your aerated butt water was pretty fully saturated with oxygen, I would be a little bit worried that the level for the tank at night was 1/2 that level.
On the basis that the fish were behaving normally, and the Otos' gill rate was nice and slow, I don't think there was an oxygenation problem in the tank before the water change.

Also, on the basis that the new water almost certainly increased the oxygenation of the tank water, yet the Otos' gill rate immediately increased markedly, I think we can rule out oxygen as being the cause.

I'm fairly convinced that the results from the kit are highly dubious!

Maybe next time I'll go and buy some RO water and use that instead of rain water, and see if there's any difference. Perhaps as Ian says there's something in the rainwater that I can't detect, and that has a temporary effect - the Otos' gill rate has now slowed down to normal, so whatever the problem was was temporary.

I must confess I've been squinting at that chart and I can't figure out how to use it! I'm probably being dense.
 
I've found the explanation of how to use the chart: you draw a straight line between the temperature on the top scale and the oxygen concentration on the bottom scale, and where it intersects the sloping line gives the percentage saturation.

What I can't find anywhere is a rough value for oxygen concentration and/or saturation below which fish will start exhibiting increased gill rate. I guess it varies from species to species, but ball-park figure?
 
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