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NanoFishNut

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  1. Sorry to break taboo and open up a 2-year old thread, but this still comes up frequently in my search results when I google comparisons between PAR meters v apps. I wonder how Photone rates nowadays. It's been under development for about 3 years now and consistently blows all other apps out of the water in regards to other tests that I've seen. For iPhone users, they usually need to make a paper diffuser using 80mg/m² (or 22 lb) paper. The paper type matters, but generally, Android users don't seem to need to fuss about with that. I was also wondering about the Apogee unit you used. Did you use the SQ-520 sensor directly plugged into your computer, or did you use a reader (SQ-500, SQ-510). The reader makes a significant difference, since the SQ-510 that we usually get for aquariums applies a 1.25 correction factor to account for increased backscattering from the higher refractive index of the water. So less light makes the transition from the water to the sensor, so it reads low. As a result, it reads 25% high when used *out of water*, since it's still applying that 1.25 conversion factor. So, when taking measurements in air, we need to divide the result by 1.25 to get an accurate reading, which would mean those apps from 2 years ago had an even higher error than you thought. We can also use the SQ-500, we just have to apply the 1.25 conversion factor ourselves when measuring underwater. Temperature has an impact too, but it generally falls <1% I have access to an SQ-510 unit, so I'm planning on making some comparisons myself in a bit
  2. Great! Divot slides can be good for some things, though sometimes the divot can exceed the focal length of the objective (particularly the 100x and sometimes the 40x). For most applications (particularly for observing algaes), a standard slide is sufficient. For larger organisms, like Cladocerans (Daphnia, etc) divot slides really shine. You can also use oil in those instead of water (or a mixture) to increase the viscosity (otherwise, a lot of such organisms are way too fast to see easily). I can't provide an ID for that image, I would honestly assume it was a bit of detritus, but the way you talk about it suggests it's a living thing? I assume it was moving? The motion might help. I haven't seen anything like that though, so I'm not much help there. I've been focusing on algae, for the most part The diversity of organisms is mind-bogglingly immense, I haven't delved too deeply into all of that. Journey to the Microcosmos is a wonderful YouTube channel that shows a lot of interesting things there
  3. Hey! Sorry, I don't get onto this forum (or any forum, really) very often, but I'm always glad to see another person taking up microscopy! It can be a really addicting hobby and aquariums give us a *ton* of fun things to look at, right in our homes. The first image doesn't have much that's identifiable, those clear things are bits of damaged cellular tissue/detritus. It can also sometimes be an artifact of plastic slides, be sure to get some proper glass slides with glass cover slips if you haven't already, they have as big of an impact on the quality of image as the quality of the microscope itself, but for a lot less money. The dark lines running through the sample look like fibers, from a cloth or something along those lines. Not many things in nature look like that, though it could also be something that's desiccated, or the detached antenna of a Cladoceran or something. The second image shows some diatoms (the brownish spike-looking things, probably Pinnularia sp., though there are tons of options) And the third image just shows some general detritus. At 1,000x you probably noticed some tiny squiggles and dots that flicker spastically... those are bacteria which really can't be identified with optical microscopy, but are interesting to note as a kind of background bustle in 800x and above You can find 3D printed darkfield and oblique lighting filters on Etsy and eBay, and you can also make them yourself pretty easily, just check out videos from Microbehunter on YouTube for info on that. Those will really increase what you can see. Over on your other thread about the algae, that Hornwort-looking thing that you saw was a green filamentous algae (looked like Oedogonium, but it was hard to see the chloroplasts, and didn't see and branching or akinetes) and it was absolutely covered in diatoms. Here's an image of Fragilaria sp. diatoms, compiled with a 13 image stack with oblique illumination at 800x (20x eyepiece with 40x objective), which I've found to be the most convenient for viewing most algaes
  4. Unfortunately, I've dealt with some of these categorizations, and I find it's better to stick with a general physical description than to guess at the identification. Well meaning misinformation is very easily spread this way. If you want to send me a sample, I'd be happy to identify it for you. It's a service I offer for free to everyone, because I enjoy microscopy as a hobby. Just let me know if you want to, since this one has been a longstanding nemesis. Algae ships quite well.
  5. I see that this is a very old thread that has been revived willingly, so I won't try to belabor this too much but: Why do you think it's Spirogyra? Without microscopy, the odds of accurately identifying Spirogyra are very low. Many species of Oedogonium (Order: Oedogoniales), Mougeotia (Order: Zygnematales), Klebsormidium (Order: Klebsormidiales), some Rhizoclonium (Order: Cladophorales), etc. look identical from a macroscopic perspective. We tend to wildly underestimate the diversity of very small things. And just as any species of plant or fish may have a shared or wildly divergent ecological niche, the same is true of algaes. Even identifying to the genus level is no guarantee that this species or homotypic synonym is going to react the same as another, let alone species from different Orders, Classes, and Phyla. In fact, your algae looks like it's not even mucilagenous, which may point more to Cladophorales. One thing we can say with some confidence is that filamentous algaes require relatively high levels of cellulose (carbon) to form their filaments. A common form of carbon that is more metabolicly advantageous to algaes than to higher order plants is organic carbon. Now, when CO₂ levels are very high (and everything else is good), plants can suppress the algae growth even in the presence of higher levels of organic carbon through whatever their algae suppression mechanism is (many have been proposed, but none have been proven). But, when CO₂ levels are not high enough (either due to poorly optimized CO₂ injection, or in a non-CO₂ injected tank) or some other factor is inhibiting plant-based algae suppression (biomass not high enough, some nutrient not high enough, etc), then the presence of moderate levels of organic carbon can be a common cause of filamentous algaes. This includes red algaes, but also includes many filamentous green algaes. So, the first thing I do when I see filamentous algaes in a non-CO₂ injected tank is clean. Clean the filters, clean the substrate, clean the plants, remove any melting leaves, do a water change after every cleaning. Then I ensure that plants are getting what they need in terms of nutrients. Low PO₄ has always caused more issues for me than high PO₄. The theory that high PO₄ causes algae has been falsified many times over in many different types of tanks, with or without CO₂ injection. That's my main complaint with Easy Green (and 90% of all commercial ferts): not enough PO₄. In fact, my number one symptom of mild PO₄ deficiency is filamentous green algae. As PO₄ deficiency get more extreme, I usually see the filamentous algaes fade to be replaced with GSA (which is primarily from the Coleochaete genus... it's much less diverse and as a result, it's much more predictable). Spirogyra sp. Another Spirogyra sp. Mougeotia sp. Klebsormidium Oedogonium We've been struggling to pin down an ID on this one, but the best guess so far is Rhizoclonium. Oddly unbranched though
  6. I'm aware of the pH pruning. As I said, the algae had only attacked a few very old leaves. For more context, this happened when I rescaped the tank over 6 months ago and nothing that had grown during that time had any signs of algae. Healthy, algae-free leaves that had grown since then also melted. Conversely, a couple of the leaves that didn't melt were algae-infested. Something else seems to be at play in my instance, but I don't know enough variables to say what. The amount of salt in Club Soda seems negligible, but since it seems to compromise comparability, I'll switch to seltzer for future experiments. I'm not sure that my cheapo microscope is up to the task of examining healthy leaves without damaging the plant, so I'll just have to be careful with Buceps in the future. For the other info I guess I'll continue wading through the thread. Thank you for all the work you've done on the subject. I'm impressed with the amount of testing and research that has gone into this, I just wish the information was compiled into a better format. It seems like the summary post has some very basic information missing and lots of overly technical detail is spread throughout, but I'm an accountant. I'm used to condensing information down into tables as much as possible, lol
  7. I haven't read through the 21 pages of comments etc, but is anyone putting together a list of plants that have been tested and the effects witnessed? I feel like such a list will be quite important, people tend to reject things that are new out of hand. If they have a negative experience early on, especially right out of the gate, then all the testing and scientific rigor in the world won't convince many people. Anticipating these issues is important. For example, Aegagropila linnaei (Marimo) is listed as "static entities" that were tested, but with almost no info on the effects on this algae. I'm also particularly interested in how mosses have been affected. I have some Fissidens fontanus that's infested with a particularly troublesome algae that I'm still trying to identify. The current contenders are Cladophora, Pithophora, Rhizoclonium, and Vaucheria. I'm suspecting, given my experiences below, that the Fiss may not tolerate RR. Personally, I've had the following results using club soda: Alternanthera reineckii var. 'Ocipus' - no issues Anubias barteri var. nana f. 'Broad White' - no issues Anubias barteri var. nana f. 'Jade' - no issues Bucephalandra sp. 'King tetana' - widespread melting, about 60% of leaves lost the next month* Bucephalandra sp. 'Mini Blue Arrogant' - widespread melting, about 80% of leaves lost the next month* Bucephalandra sp. 'Blue Sekadau' - widespread melting, about 80% of leaves lost over the next month* Ludwigia repens var. 'Some Ditch Behind My House' - turned pale and leaves wilted, new growth is fine** * None of these Buce were new arrivals and I've moved them in the past with no melting. The fertilization, light, and CO₂ all remained constant. The only variable that changed was the RR. They just a tiny bit of BBA (Andouinella), Staghorn (Compsopogon), and GSA (Choleochaete) that built up a long time ago and had been stalled for quite a while. I thought it might be nice if it cleans up the older leaves rather than being forced to trim them. Instead, I've lost almost all of the leaves, including leaves that had no algae. **The Ludwigia isn't a valid test of RR because I dosed H₂O₂ first, then later decided to do a round of RR. Thus, the result is likely influenced by the H₂O₂. The tests that I've seen so far have all been on common, hardy plants in low-tech setups. I have 40 species of plants just in my 75 gal alone, many of which are rare, finicky plants, in a high-light, high-CO₂, aquasoil, and rich fertilization, Garden Style environment. My tanks also run highly acidic, which could theoretically affect how RR works. My typical pH levels in my tanks are 5.8-6.2 before CO₂, and 4.8-5.0 during the day with CO₂. Once my busy season is over with work I have some plans to perform controlled testing with trimmings of my plants. My plan is to set a couple stems of each plant into two containers, one with tapwater, one for RR, together in the same location (so the same light, etc), treat them the same, and then replant them in the same tank they came from to see if there's any difference in how they do. I won't be able to test this with ever plant that I have (there's no way I'm gonna trim my Hygrophila araguaia f. 'Chai" for this, or subject my rarer Buceps to such treatment). It would be nice if there was a location I could post the results so they could be compiled into a single location. I doubt I'll have the time to do it myself.
  8. Mmm, that's not A. crispus in the front left. I see what might be a crispus leaf in the back right-ish though. A. crispus has pointed leaves, the front left looks like A. ulvaceus, though it's worth noting that most, if not all, Aponogetons in the hobby are heavily hybridized. Your Anubias has been experiencing a long-running phosphate deficiency. I've personally never seen GSA without phosphate deficiencies. Common with Easy Green, it's quite low in phosphorus. Leave it long enough and the rhizome will decay, that's the plant consuming its storage because it's not getting what it needs from the water column.
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