Forum Replies Created
-
Posts
-
[quote=”An_Outlier” post=5409]Do any of the more experienced people here… [/quote]
I don’t count myself in that number. Nevertheless I may be able to contribute to the topic (for which, thanks – because it’s been on my mind from time-to-time, too).
I have three or four electronic pH-measuring devices : a hand-held ; and a couple of in-tank monitors (but not in Paros tanks). A noticeable feature is that, when freshly cleaned, calibrated and challenged with identical samples, they all return slightly different results! In context, the differentials are perhaps not significant but they’re there nonetheless.
I trust my hand-held one more than the others and use it for Paros water. It’s one of these. I’ve been using it for a couple of years and it hasn’t yet told me that it needs a new electrode. There’s another version (pH110 I think) that has a refillable electrode and which I may go for when the time comes. In the meantime, I’ve found the pH100 really easy to use and to calibrate. I use an Extech conductivity meter, too, and am similarly pleased with it.
One difficulty we have is that electronically testing for pH in low-conductivity water may be somewhat unreliable (because of the way such meters work) so the results must be viewed as indicative rather than absolute. Still, I would rather have some idea than no idea of that parameter!
I guess that the more ‘difficult’ the species of Paros, the greater concern an optimal pH (and/or conductivity) would be.
If I were to abandon my meter and rely on intuition and oak/catappa leaves, I would have to schedule myself to renew the leaves regularly : after a few weeks’ immersion (I found in tests) they cease to acidify and the pH of their host water may actually increase.
Over to the experts : I look forward very much to reading what they have to say….
[Incidentally, does your Username indicate that you’re a Geologist?]
re: getting rid of Hydra (just in case this info is useful) …
I had green Hydra in my Hyallela tank and can report that I got rid of them using NT Labs Anti-Fluke and Wormer. The active ingredient is flubendazole, as in many other antihelmintic preparations. This one is convenient, though, because the flubendazole is pre-dissolved (it dissolves extremely reluctantly in water).
I rescued as many Hyallela as I could and parked them in temporary accommodation while I treated their tank. The day after the first dose I inspected the tank. At first it appeared that all the Hydra had gone but I did eventually find three that seemed still operational (I poked them with a skewer and they collapsed!). So I applied a second dose, after which I haven’t seen any Hydra at all in that tank. Incidentally, the Hyallela that remained in the tank during treatment all survived it.
I now have to deal with an outbreak of Hydra in my blackworm tank, which might be more problematic!
I’m glad that my link worked!
Bennie : all my filters have been seeded, chainwise, from my original Discus tank. Although that tank was decommissioned nearly fifteen years ago, all subsequent tanks (those that contained fish, anyway) have been of the soft/acidic variety, so I would expect that the original assembly of micro-organisms would largely have duplicated through the chain. Whether or not there was a mixture of bacteria and archea, it seems that archaea were at least dominant : that appears to explain why the filters’ nitrifying micro-organisms behaved in ways that bacteria simply could not have done.
Without rummaging through my notes I have it in my head that the nitrifying pathways of AOB and AOA differ : the former extract their carbon from carbonate ; and the latter from dissolved carbon dioxide. The more acidic the environment, the greater is the fraction of dissolved carbon dioxide present (I think I’m right in saying that), hence one reason why such environments tend to select for AOA.
I thought I’d jump in here with some personal observations resulting from a (very!) amateur experiment. I may well have mentioned it previously in this forum and, if so, I apologise for the repetition. Nevertheless, the appearance of this thread allows me the opportunity to ‘bump’ it and perhaps to bring it to the attention of new readers. So …
In the Spring of 2011 I found myself with a 12-litre tank which had previously housed some shrimp and a few baby Ancistrus. They had all been evicted some four or five months before and I had kept this little tank ‘running’ in the interim with its heater and air-driven sponge filter. It was unlit apart from ambient light. The only visible signs of life had been some moss and a few Malaysian Trumpet Snails. Other than topping up with water to counter evaporation loss, I had paid the tank no attention at all.
Eventually the time came when I needed to use the tank (I noted that its pH was 8.04 at this point). Given all the facts about aquarium nitrification that I had come to understand from an appreciable amount of reading of authoritative sources, the filter surely must have ‘died’. So I set about recycling it using my favoured variation of the ‘fishless cycle’ method (it involves adding both ammonia and nitrite). To my astonishment, the filter began working immediately. I have just found my notes from the time – I see it was in March 2011 – and, at the risk of boring you somewhat, can extract some detail from them.
I read there that I had decided to test to see if the filter would make any effort at nitrification and, further, that I would start by adding nitrite alone. I guessed (rather generously) at the amount of nitrite that the assembly of previous occupants might have produced via ammonia oxidation. As a result of that calculation, I added nitrite to an in-tank concentration of approximately 0.9mg/l at 1330hrs on 27th March 2011.
I hadn’t expected that anything would happen and at that stage I thought I might test after 24 hours or so. However, for some reason I tested at 1630 the same day : the in-tank concentration of nitrite measured approximately 0.4mg/l. By 0130, i.e. 12 hours after inoculation, nitrite was undetectable. I repeated the test with a greater concentration of nitrite and this, too, was oxidised in quick time.
I then ‘reset’ the tank with a large water-change and tested for any ammonia-oxidising activity : I used the Seachem test kit, which allows both unionised and ionised ammonia to be estimated. I began with an in-tank concentration of approximately 0.31mg/l ; unionised ammonia was undetectable nine hours later. I followed this by adding ammonia to an in-tank concentration of 4mg/l ; a zero for unionised ammonia was achieved somewhere between 38 and 49 hours later.
I subsequently carried out other ammonia/nitrite procedures on this tank but even at this early stage my suspicions were aroused that all I had learned so far about nitrifying bacteria was not necessarily true : without ‘food’ (so the texts had said) the bacteria would die off quickly – some sources said that this would happen in a matter of hours.
To double-check I removed the filter media, packed it in a bottle of fresh water and posted it to someone who had volunteered to help out, in Yorkshire. Her instructions were to remove the cap from the bottle, keep it unmolested for a few days and then send it back to me. In the meantime I had once again ‘reset’ the filterless tank with water-changes. When the filter media arrived back, I reinstalled it in the tank and repeated the ammonia/nitrite dosing and testing : there was no significant difference compared with the results achieved prior to the filter media’s ‘holiday’. [In retrospect I should have sterilised the tank in the absence of its media ; but even without sterilisation, I reason that I should have been able to detect some difference in nitrification ability]
I had read claims from some hobbyists that nitrifying bacteria are capable of dormancy ; and from rather more trustworthy sources that nitrifying bacteria are incapable of encysting and therefore are incapable of dormancy. If the latter were correct (I thought) then agents other than nitrosomonas/nictrobacter-type organisms must be responsible for nitrification in my little tank.
I needed to use the tank but wanted to keep the filter media as it was. So I moved it to a 15-litre bucket which I equipped with a heater and airline. Over the year (and more) that followed I subjected the media to a wide range of environments. I played about with inter alia, with temperature, pH, ammonia/nitrite concentrations, mineral concentrations etc. : its nitrification capabilities remained. The only condition I found that would reliably shut it down was when pH fell to around 3.4 (as measured by my meter – I know that this could be an indicative, rather than an absolute value!). To summarise : it became absolutely clear that whatever was doing the nitrification wasn’t behaving like the bacteria that authoritative authors had been writing about, inclusive of the admirable Tim Hovanec.
I was both baffled and intrigued ; I carried on through the Summer reading all that I could find and vaguely understand (I’m not a scientist!) about nitrifying bacteria, but nothing surfaced that helped . Then in October, idly surfing the ‘net using my set of keywords, I happened upon a paper that appeared to offer an explanation. It was this one (if the link doesn’t work, search under its title: “Aquarium Nitrification Revisited”. You may have to click on ‘Article’ if the abstract or full text doesn’t appear straight away).
Briefly : a Canadian team had been looking for environments in which to study archaea. Among the environments considered were aquaria, both freshwater and marine. Using Polymerase Chain Reaction analysis they were able to determine that in a significant proportion of the aquaria studied, genetic components from ammonia-oxidising archaea (AOA) far outnumbered those from ammonia-oxidising bacteria (AOB). In some aquaria, no ammonia-oxidising bacteria seemed to be present at all. Conditions which select for AOA include low nutrient concentration (i.e. low ammonia) and low pH.
Eventually, I plucked up the courage to email one of the authors (Josh Neufeld) and outlined what I had been doing. He agreed that my observations were consistent with nitrification by archaea in my tank/bucket. As an aside, I followed up with a tsunami of questions to Dr. Neufeld about the nature and behaviour of AOA, looking to compare and contrast with AOB : from memory, of my twenty questions he was able to answer just one of them with certainty, so recent was the concept of AOA at the time. Maybe more is known now – if anyone reading this follows it up and discovers (or already knows of) new and relevant information, I’d love to hear about it.
Sorry about the length of the above but I hope it has been of interest.
(I considered adding this to Jacob’s ‘opallios breeding’ thread, but in the end didn’t. I don’t think it deserves its own thread, so I’m tacking it onto one of my old ones! Please move if necessary)
A while ago I moved a pair of opallios into a 15-litre compartment in a row of similar tanks that share the same water. It had been an unusually long while since I last saw either, so at this week’s water-change I thought I’d have a hunt for them.
I began removing wood, moss etc. starting at the front of the compartment. Within a few seconds the female appeared, shocked and disillusioned from underneath a pile of leaf-litter. I continued the search for the male …
Lifting up a bamboo cave I came across what I presume to be a bubblenest (pictured here) :
I say “presume to be” because, although previously I’ve had several spawnings of opallios, I have never noticed a bubblenest before : eggs were ‘bare’ and simply stuck onto the roof of a coal cave.
Was I therefore previously unobservant ; or do opallios in fact sometimes use bubblenests and sometimes not? I can’t see any eggs in this bubblenest, incidentally. Having taken the photo, I immediately put everything back as close as I could to where it was before. Even so, I assume that I’ve ruined any potential of this particular nest.
The question that follows on from this is : would I be justified in resuming the search for the male in due course, or can I take it that the presence of a bubblenest is sufficient evidence that he’s lurking there somewhere?! (EDIT : it’s OK – when I returned downstairs there he was, stationed at the front of the tank wondering what on earth had happened!).
re: Catappa leaves : an alternative source to consider …
I get mine from Jeff & Wan. Whilst buying a batch of ‘perfect’ leaves from them, I also get a bag of chopped-up low-grade leaves with which I make ‘blackwater extract’. However, having now sourced Aquahum as a result of a mention by (I think) Pavel, I may just use that instead.
I have done similar (i.e. have a relatively-larger tank subdivided) and attach a piccie of the assembly. The sponge plugs between the tanks are c3cm thick : I can’t imagine much getting through that would compromise the integrity of (say) two different species in adjoining compartments …
… except that I have lingering worries over particularly enthusiastic sperm! This is because there is a current, albeit very slow, from the left-most compartment to the right-most.
Is the possibility of the transmission of sperm from one compartment to the next something that I need to keep in mind, do you think? Or are they so short-lived that it’s not an issue?
Just an update …
I’ve now found a source for Dero Worms ; it’s actually the same place as my Moina came from (the USA) – but it’s a relatively recent addition to their catalogue.
If my maths is correct (mostly it isn’t!) then 0.15mm can be expressed as 150 microns.
There are sets of artemia sieves that you can buy. The set that I have has the brand name “Hobby” and consists of four sieves (5 x 5cm) of 900, 560, 300 and 180 micron mesh. I think JBL offer similar sets – though I don’t know their mesh gauges.
I also have a 53 micron sieve bought from here. If you scroll down the page you’ll come to it ; and you’ll also see a 125 micron sieve. Alternatively, you can buy the mesh separately and make your own sieves!
There surely must be an equivalent source in your country : the keywords to use for a search are the appropriate translations of ‘mesh’ and ‘micron’.
If not, then I’m sure ZM Systems will send overseas.
My own Moina culture method has been OK but results in a variable supply. Numbers will build until there are absolutely zillions of them, then recede until there are only a few hundred or so before building again! I’d like to achieve a more dependable supply and so I’m going to try your method, Deepin Peat. To that end I’m making myself a guide based on the information that you have kindly offered in this thread.
One question : presumably a culture based upon your method doesn’t continue for ever? So is there an optimum duration that you’d allow a culture to run ; and how do you tell if a culture requires renewal ‘early’, as it were?
I haven’t the faintest clue what the rationale for this statement could be. I’m intrigued, though. It’s a pity my German’s not up to it, else I’d ask him! His e-mail address appears to be: guenter-kopic@t-online.de
Do you (or anyone) feel like giving it a go?!
Is this fresh peat or is it dried loose peat that you are rehydrating? If the latter, I have some notes somewhere that I’ll try to dig out for you in due course. If the former, then so far as the acidifying effect is concerned, it starts its work straight away.
The turbidity you observe is from very small particulate matter : I’ve played with it and it passed through a 53-micron screen! A few thicknesses of kitchen paper gets rid of it (though it’s very slow and hardly convenient for in-tank use!). There’s less turbidity from fresh peat than there typically is from rehydrated. I don’t know about compressed dried peat (pellets, or whatever) because I’ve never played with it.
Other than its effect on pH, peat leaches humic acids into the water column. If it’s anything like leaf-litter (oak, catappa etc.) tannin is leached first ; then as immersion continues, fulvic acids. But I don’t know whether peat does that – it must be loaded with fulvic acids, so I guess the fish must get some benefit from it right away?
Incidentally, I’m playing with some fresh peat at the moment. I was startled to note the presence of nitrite. (It was at a low concentration but, as we know, nitrite toxicity to fish increases alarmingly with decreasing pH). Have you been able to observe something similar with the peat you’re using?
You can use dipslides to roughly ascertain the numbers of aerobic bacteria in your water. The type you’d need is BT2 – they’re easily available and are less than £10 for a box of 10 in the UK.
Update …
At least two of those eggs that were carried downstairs, stuck on a piece of foam, have hatched!
Yesterday I thought I’d take a pair of opallios out of their current accommodation and give them a room of their own. The most suitable place was what was my shrimp tank, so I evicted all the shrimp into a third tank and did a thorough Spring clean of that ex-shrimp tank. The whole process required logistical ingenuity ; it took several hours and there were containers full of water everywhere!
Before moving any Paros, it was necessary to take everything out of the tank ; and before doing that I had to make sure that no fish were lurking in the caves [you may like to refer up-thread to make sense of this!] and prevent them going back in before the water-level dropped. To do this, the rectangle of foam that blocks each window outside the tank is removed and transferred inside the tank, blocking the entrances to the caves. So far so good.
I found a Paros fry that I didn’t know I had ; and managed to transfer two adults (one’s definitely a male and I hope the other is female) to their new accommodation.
After a lot of faffing I was at last ready to reconstruct the main Paros tank. I began removing the foam rectangles from the caves … and was astonished to find a load of eggs adhering to one of them (it was the one picture in Reply #2415, in fact). A quick glance in the cave told me that I hadn’t been very attentive when blocking the cave earlier : there was a male in there – and he must have been trapped in there for about four hours!
I couldn’t immediately think of a way of re-introducing the twenty (or so) eggs to the cave, so I scattered them in a clump of moss in the ex-shrimp tank. They were extremely sticky, but I got them all off the foam eventually. I don’t know what the likelihood is that they’ll hatch? Apart from the pair of adult Paros, the only creatures I deliberately put into that tank are a few cherry shrimps and a small gang of Asellus aquaticus ; I’m sure the Asellus won’t touch them, but I guess the shrimp may find them too hard to resist. I guess I’ll find out in due course!
Back at the main Paros tank, I inspected the cave more closely. There are still around twenty eggs stuck to the roof. During the process of removing everything from the tank, hovering and then doing a partial water-change, the water-level had dropped such that it had been a millimetre or two below the eggs : they wouldn’t have dried out as such, but they would have been exposed to air for at least some of the time. The male is still in there guarding them. That may be a sign that all is still well ; or it may be a sign that he is completely stupid. I guess I’ll find that out in dies course as well!
A couple of bad piccies. The first shows a bit of the male in the cave (before the camera flash made him scarper!).
The second is a fish’s-eye view of the cave entrance, looking out into the tank.
Although I can see the remaining eggs, I’m afraid I can’t get a photograph of them at the moment.
