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This is an area of particular interest to me. Perhaps I may be able to share some bits of info that, while they may not answer every question, may at least provide food for thought. I should add that I am an intrigued hobbyist ; I am by no means a trained scientist!
All my tanks are filled with soft/acidic water. I had become casually interested in nitrification, reading as much as I could in the fishkeeping context and a little wider. The received wisdom was that agents for nitrification in fishtanks are bacteria – such as the oft-quoted nitrosomonas/nitrobacter team. This belief stemmed from studies of nitrification in an industrial context (sewerage treatment etc.) and spawned various preparations that appeared on fish shop shelves … and didn’t work very well.
More progress was made by Tim Hovanec, who proposed that different genera of bacteria were responsible for nitrification in fishtanks. A new generation of ‘Bacteria-in-Bottles’ (BiB’) appeared in shops based on his research while he was employed at MarineLabs. They were more successful ; Tetra Safestart and its Americam equivalent, for example, are ones which are frequently mentioned in forums as having at least a chance of working, given the correct handling. (Tim Hovanec later left MarineLabs and began Dr. Tim’s Aquatics, which now markets its own BiB’s).
‘Bactinettes’ was another bacteria-based initiative which failed because shops couldn’t maintain the conditions necessary to keep the live cultures viable (even though the company provided refrigerators for the purpose!).
However, my experience of looking as closely as I could at nitrification in my own tanks seemed curiously different in detail from how it should have been if bacteria were indeed at work. I remained puzzled until …
… in October 2011 as I was surfing the web using relevant keywords. I came across a very recently-published account(1) from a Canadian group who had been evaluating various environments in which they might be able to study archea. One of the types environments that they looked at, knowing that ammonia would be generated in them, was hobbyist fishtanks. Analysing RNA, they found that the main (sometimes the only) nitrification agents present were Ammonia Oxidising Archaea rather than Ammonia Oxidising Bacteria – even when BiB’s had been used to begin a tank’s cycle. My experience began to make a little sense at last!
There have since been further papers produced by the same lab that suggest : the more acidic the environment and/or the lower the concentration of nutrient (i.e. ammonia) the less likely it seems that bacteria will be the main oxidisers.
If we rely on the hobby’s still-current wisdom, then indeed : nitrification should stop completely given certain conditions. Amongst these conditions would be, for example: a pH of <6 ; or an absence of carbonate. However during the past couple of months I have been running a bucket containing an air-driven sponge filter which has been oxidising to completion c1mg/l ammonia at pH5 and at a KH that doesn't register on my Salifert test kit (the lowest it measures is 0.3mg/l). Admittedly it takes nearly 36 hours to process each dose (it's operating at room temperature) but I think you'll agree that this rather flies against the hobby's belief in nitrifying bacteria!
A further point that seems relevant to this thread (though I'm absolutely sure that some/most/all of us must be aware of it already!) is the equilibrium between unionised (or 'free') ammonia (NH3) and ionised ammonia (NH4+). To explain, just in case you’re not confident about this …
When ammonia is bubbled into water it dissolves very eagerly. As it does so, a proportion of it becomes aqueous NH3 and a further proportion of it becomes aqueous NH4+. The relative amounts that are formed depend upon certain properties of the water : temperature ; pH ; and salinity. The two more influential properties are temperature and pH : the lower either or both of these values, the greater will be the proportion of the relatively harmless ionised ammonia present.
Most of the test kits that are available to us report on the concentration of Total Ammonia present – that is: the sum of both ionised and unionised ammonia. The kit (I think) that you’re referring to, Jennifer, that measures each separately, is Seachem’s Multitest. Interesting as it would be for you to have one, it may not be necessary to lighten your purse in order to find out how much of the toxic unionised ammonia you have in your tank(s)! The relative concentrations can be derived from a calculator(2) provided that (as is likely) you are able to make reasonably accurate determinations of temperature, pH and conductivity.
Let’s imagine an unlikely scenario where you’re panicking because your salicylate ammonia test kit (you can’t use a Nessler kit in a typical Paros tank, of course!) reports Total Ammonia at 0.25mg/l. We’ll assume your tank is at vaguely-typical values : temp 26C ; pH5.2 ; conductivity 60uS/cm. The calculator declares that the toxic ammonia in it must be 2.33E-05mg/l. Now, I have no idea what that means (I’m pretty much innumerate!) but I know it’s a very small number. I have to crank up the Total Ammonia concentration to 1.5mg/l before I see a number that I recognise : 0.00014mg/l of toxic ammonia, which is still insignificant for practical purposes.
Total Ammonia generation in a Paros tank will be extremely low, if only because of the light stocking and the comparative low activity of the fish. There will be some nitrification being done by archaea (it should be a little faster in a tank than in my bucket because of the elevated temperature) and some ammoniacs will be used by plants (if they’re there). So, in theory, water-changes shouldn’t be required as frequently as they are in (say) a community tank filled with tapwater. Water-changes still need to be done of course (in my opinion) in order to maintain the redox balance in the tank.
I hope at least some of the above has been of interest and/or value!
{Incidentally, for the first time I noticed one of my males ‘displaying’ briefly a couple of days ago ; it was jaw-droppingly beautiful. I hope I’ll be able to get a decent photograph one day …}
1. Aquarium Nitrification Revisited
2. Fish Hatchery Management (Table 9)[quote=”Little” post=2426]Thanks Helene — that worked. We were looking for #1146. Jalmj you are looking for Vale! (Mr. Love) and you are in luck he is in the UK (Milton Keynes, UK). He was able to order moina culture previously in the UK. Hopes this helps.[/quote]
Hey – that’s me!
Coincidentally I’m just about to order another culture (my previous one having collapsed. I’m still unsure as to whether the source is happy to be made ‘public’ … though maybe I said who it is in the post(s) that Bill (et al.) refers to? If not, then please message me, jalmj, and I shall give you the relevant info.
Cheers
Perhaps this piccie of a newly-constructed cave will help?
I then superglued moss to them.
Incidentally, I’ve seen another fry, much bigger then the one I spotted earlier – so it seems possible that there have been two clutches of eggs laid. I haven’t been able to get a decent photo of it (it’s genetically programmed to be aware of the distance it must be from a camera in order not to be photographed!)but this is the best I can do for the moment :
When I put in a second helping of green water, some mosquito larvae were included. I watched as this fry considered one for a long time. Given the relative sizes of fish and prey, I thought there’d be no way that the former could fit the latter in its mouth. Wrong!
Just a quick update (now that I’ve remembered to post one at last!) …
One single larva hatched from the frozen rafts : not enough of an encouragement to pursue that particular storage method!
Yesterday I put a few rafts in the freezer, just to see what would happen. I’m about to take them out : if there are signs of life at some point, I’ll let you know!
I can confirm that zillions of tiny larvae have exited from the egg-rafts (had to use a magnifying-glass, though!).
So hatching occurs in less than 24hrs at c30C.
A bunch of rafts have appeared in one of the outside tubs that I use as live food sources. They weren’t there ysterday evening. I’ve collected some and have put them in a separate, small, container filled with their original water. I’ll observe and let you know when they start to hatch : it’s very hot here at the moment (c30C) so presumably hatching will be accelerated?
My normal regime is a c50% change every two weeks. However …
… I have just, coincidentally, finished a period of heroic restraint during which I did nothing to my tank (except add food) for seven weeks. I wanted to see what would happen.
I’ll be writing up the results in due course. I’ll be posting them in the general fishkeeping forum that I inhabit, from whose Members there’s been some interest because of the Paros’ (relatively) extreme conditions. I’ll post a link in this thread when I’ve done it in case they’re of interest to anyone here and hoping that you may offer some helpful criticism . It’ll be some time yet, because I want to try to establish the reasons for one or two of the results ; and I’m going to wait until the tank has been well-settled at its former parameters before repeating both a dip-slide test and an assessment of where some micro-organisms seem to be concentrated.
Well I would gladly live in Wellington (New Zealand) if given the opportunity. However, someone has to live in Milton Keynes (UK) – and I drew the short straw!
No – my comment was in reference to my following a cricket match that was ended by falling water!
Hi Helene
Yes – I’ve now seen your video and I think you’re more likely right with Paramecia (or something like them) rather than Ostracods. Your critters seem to be much more at the mercy of the tank’s current and occupy the whole water column ; while mine prefer to associate themselves with surfaces in their tank – although they do zip around when they feel like it!
Since it’s raining in Wellington (New Zealand) I did you a quick video of my Ostracods anyway! But probably nothing you hadn’t seen already during your earlier Googling. Here goes attempting the link …
I have ‘clouds’ of similar-sounding creatures in one of my shrimp tanks. They are Ostracods.
Would you like me to put one under a microscope so you can see what they’re like?
A Brit’s-eye view …!
A typical torch was/is a bundle of burning oil- or wax-soaked rags tied to a pole (enabling it to be portable) and set alight to be used for constant (if a bit smelly and smokey!) illumination. The modern portable equivalent, using batteries and an incandescent bulb, is therefore aptly described as a ‘torch’.
However, the battery-powered version also affords switching on and off the illumination in rapid succession – useful for signalling in Morse Code, say. So its description as ‘flashlight’ is equally as valid, irrespective of the duration of a ‘flash’.
The advent of LED versions with tiny round batteries, of course, means that they don’t even have to be torch-shaped any more, though the other properties remain!
‘Taschenlampe’? I really like that word – it sounds to English ears as though it defines a lantern powered by sneezing!
[quote=”Jordy” post=1700]PH 5,5 … KH 0 … Just want to cycle the tank [/quote]
Genuine question (because I have a special interest in this area):
What do you expect to happen, cycling-wise, with these parameters (which is close to what they’d be when you stop ‘cutting’ your rainwater with tap)?
And, with plants in the tank, what measurements will you perform that will leave you in no doubt that cycling has taken place?
