Dying Caulastrea Coral Rehabilitation
Coral
Dying Blastomussa Coral Rehabilitation
Dying Australophyllia Rehabilitation
Dying Acropora Coral Rehabilitation
Under construction! (2012-ACRO001)
Turn it up to 11: Magnetic Stirrer Coral Dip Station
Sure, blowing a coral with a pipette while it sits for 10+ minutes in a dip works. So does blasting it with a powerhead. But… I’m a nerd, and nerds must always over-engineer every problem (or non-problem). This project was the result of trying to find the end of the internet one night while researching new 3D printing ideas. I was inspired by ChickenHaunt’s Resin Wash Station over at Thingiverse. With only a minor tweak, I now have an amazing Coral Wash Station! Enjoy “turning it up to 11” with this magnetic stirrer.
Supplies List:
This is a list of what I actually used to build my magnetic stirrer coral dip station, along with estimated prices as of the published date. I’ve included links to the products and 3D print files (please note, as an Amazon Associate I earn from qualifying purchases. I get commissions for purchases made through links in this post.) While other stirrers and containers will still make a nice coral dip station, they may not work with the 3D print file. Therefore, I highly recommend this list since it is all most likely to work well together. I also recommend fluorescent filament for the print file, since the setup will likely be near a reef aquarium (so the filament will glow!) At the end of this post, I’ve included more details about the supplies, along with some other options.
- Intllab Magnetic Stirrer: ~$30 USD
- 3D-printed Rack with fluorescent filament: ~$3 USD, if printing yourself
- Magnetic Rods with Retrieval Rod: ~$17 USD
- Sistema KLIP IT Lunch Plus Storage Containers: ~$9 USD (recommend getting two sets – or more if you do a lot of dipping)
Coral Dip Station Instructions:
Step 1: Prepare the Magnetic Stirrer
First, I started with the Intllab Magnetic Stirrer. This is a simple stirrer that does not have a heating option (not necessary for how short coral dips are). Now, if you wanted a coral bath option, I recommend considering a heated magnetic stirrer (but the cost is much greater).
One of the most important qualities of this stirrer is the 316 stainless steel plate due to the corrosive nature of saltwater. (Note: this metal is corrosion-resistant against saltwater, not corrosion-proof. Try to keep the stirrer as clean as possible for the longest life.)
Step 2: Prepare for 3D Printing
My 3D printer is a Creality Ender 3 Pro, which is a “best value” printer in my opinion. The fluorescent green filament I used for the stirrer tank surround is in the storage tub on the left. The black filament used for the rack is on the right.
Step 3: Print and Build
I followed ChickenHaunt’s instructions for the printing and assembly, but there are a few remixes out there on Thingiverse too. Here are a few things I liked/didn’t like along with tips:
- – Use solvent weld to assemble everything rather than hot glue and/or super glue
- – Assemble the components on the stirrer before gluing; otherwise they may not fit (including the magnetic stirrer and the storage container)
- – The strainer platform is quite flimsy
I added a sticker with the infamous quote, “Turn it up to 11!” from the movie, Spinal Tap. There’s no “11” on here, but most of the time I turn the stirrer up to the 4th blue bar. Use trial and error to see what works best for your corals and setup.
Step 4: Add Parts
Once printing and assembly is complete, add the dip container. It should fit snugly into the surround. Of course, the larger containers that ChickenHaunt used would work great for larger corals (but those containers can be difficult to find. I like these smaller ones for frags and small colonies.) Add a magnetic stirrer rod.
Note: The magnetic stirrer kit I recommended includes a retrieval rod. Personally, I try to keep my hands out of coral dip as much as possible for safety. The retrieval rod will pick up the magnetic stirrer rod, thus helping to prevent skin contact with the coral dip. There are additional options at the end of the post.
Now, add the strainer platform with handle. The platform is very flimsy, so whenever I lift out corals, I have to support the handle’s opposite corner. A platform remix would likely fix this, but this still works fine.
Step 5: Test it Out!
Now, just add tank water and medication of choice. Plug in and turn on the magnetic stirrer. I usually turn the knob to the fourth blue bar. Once the medication is adequately dispersed, add in the corals and set a timer for the appropriate dip length.
No more pipette blowing or giant powerhead necessary!
Inspect the corals for damage and health (if you need a primer, check out Assessing Coral Health).
About a minute before the dip time is up, I turn off the magnetic stirrer. This allows the gunk to settle to the bottom. Once the time is up, carefully lift the platform out of the solution (using additional support as necessary).
I have six of these storage containers, as I typically have a three-dip process. Then, I use a fourth container for plain tank water to rinse the corals.
Magnetic Stirrer Coral Dip Station Conclusion:
When I first built this magnetic stirrer coral dip station, of course I was excited. But I had no idea how much it would transform my dip successes. The small footprint combined with the fast water movement made dips easier and more effective.
Additional Supplies Info:
NOTE: As an Amazon Associate I earn from qualifying purchases. I get commissions for purchases made through links in this post.
INTLLAB Magnetic Stirrer w/ stir bar set & retriever
INTLLAB Magnetic Stirrer
Fluorescent PLA filament
Magnetic Stirrer Mixer Stir Bar and Retriever
Sistema KLIP IT Lunch Plus Storage Containers
Creality Ender 3 Pro 3D Printer
Coral Dip
Blue Sun Coral (Tubastraea sp.): A Natural Rarity?
When I started in the hobby in 2008, there was a legitimate problem with dyed corals and anemones. Sadly, this was not a new problem at that time either. Bleached corals (or even healthy corals) were subjected to dye baths in a dizzying variety of colors, especially in rare shades, such as yellow and blue. Unfortunately, that practice had unsurprisingly deleterious effects on the animals, as the dye interfered with their photosynthesis via zooxanthallae. Fast forward over a decade, and the frequency of dyed corals showing up in local fish stores for sale substantially decreased. So, imagine my shock when a fellow reefkeeper sent me a message about a potentially dyed blue sun coral. The earliest reporting of blue sun corals in the hobby I could find dated back to 2017.
Panic Hits – Artificially Dyed Corals in a Pandemic:
In June 2020, right in the middle of a global pandemic, I was appalled to see reports popping up all over multiple reefkeeping forums and other social media sites claiming that there was a sudden influx of dyed corals. Rehabilitating dyed corals is no picnic; it takes daily feedings, lots of carbon to absorb any free dye, and fast removal of necrotic tissue. Even then, I have never had success with them long-term. Panic started to hit me, as my aquariums were severely neglected thanks to trying to telework and homeschool young children simultaneously. Not only were my tanks ill-prepared, but I couldn’t get the supplies necessary for rehabilitating dying coral. Chlorine bleach and hydrogen peroxide are two of my main staples to keep clean equipment, and they were nonexistent on shelves at the time.
Assessing Blue Sun Coral Health:
Then, on 8 July 2020, a fellow local reefkeeper posted on Facebook that she had just received one of these (on 18 June) and suspected it was dyed and dying. She was concerned that it was affecting her yellow sun coral too, as it seemed to be losing color. So, it was time for me to do some research!
Although I had a lot of questions, my first question was on how to stabilize the coral. Since the owner believed it was dyed, I needed to determine what dyes could have been used so I could figure out how to remove it. Sun corals are often various shades of orange or yellow, so the idea of getting a solid blue coral seemed a bit crazy to me. In other words, I thought that just adding blue dye to a yellow coral would likely result in a greenish coral. However, I’ve used methylene blue extensively, and I know its staining power! It seemed a plausible dye to overcome the yellow.
While researching the various dyes, I found research suggesting that anemones could safely tolerate methylene blue dye, as it did not affect the anemones’ growth or survival. Additionally, the research found that the anemones only retained the methylene blue for approximately six weeks. This gave me hope. Maybe the sun coral was dyed with methylene blue, so it might live if I could feed it adequately (as a non-photosynthetic coral, there is no zooxanthellae interference). I was also prepared with activated carbon to absorb any leaching dye (although this research on using dragonfruit skin was super interesting!)
Bringing the Coral Home:
Only three days later, on 11 July, we met in a mall parking lot, while wearing masks and maintaining social distancing. Even in the hot mid-day sun, the coral was a shocking blue color. I quickly took both the blue and yellow corals home. I followed my standard coral assessment procedure, and several things concerned me.
Determining the Coral’s Condition:
First, I’ve never previously had my hands or equipment get any sort of coloration on them while fragging. Was this the “dye?” I could see blue, yellow, and green colors on this while towel.
Second, the yellow sun coral had a case of “brown jelly disease” starting. This was not good.
Although the tissue appeared dyed, the skeleton of the coral was white as usual. This was not what I expected, especially if the coral was submerged in dye. However, the anemone research referenced above suggests a dye method that may work on just the tissues. At this point, I wasn’t convinced either way on if it was dyed.
Quarantining the Blue Sun Coral:
After initial treatments, I placed the corals into my quarantine tank for observation. I started them on small pellet food, as sometimes that seems easy for corals to digest. Once the coral was regularly eating, I switched it to a varied diet of Roti-Feast, Oyster-Feast, Phyto-Feast, Reef-Roids, and baby mysis shrimp.
Unfortunately, the “brown jelly” spread to my other corals in quarantine, but I was able to keep it from progressing on the yellow sun coral or to the blue sun coral.
About a week after arrival, I was once again convinced that this coral was dyed, as the healing fragmented portion was a dark greenish-yellow. Ah ha!
But this coral decided to just keep me guessing. Wouldn’t the tentacles be blue as well if it was dyed?
Blue Sun Coral Grow-out:
By a month after arrival, I was sure it was natural, as it was just becoming more blue with bright yellowish green polyps. How was that possible?! Methylene blue should’ve mostly dissipated by that point. It had to be natural, right??
Oh but wait… This coral just wants to keep us on the edge of our seats! After four months in my tank, the shadowed areas were turning yellowish green! But, notice the baby polyp – it’s blue! My head was starting to hurt. (And yes, unfortunately I was not feeding the coral as much as necessary, so it was receding. My tanks were quite neglected during COVID-19, as you’ll see by the algae and aiptasia.)
In reviewing the 2017 post photo, the shaded regions display similar coloration.
Spawning of the Blue Sun Coral!
On 12 December 2020, five months after getting the coral, I woke up to find this blue sun coral spawning! And, the planulae were orange (not blue) – suggesting the coloration was natural (and reproduction hopefully not affected).
One of the planulae immediately settled onto the glass, where I watched it eagerly (until it disappeared about three days later).
Here are some of the planulae. Unfortunately, none of them survived, or maybe they weren’t even fertilized in the first place.
According to username, “Tennyson,” on Nano-Reef, the dark orange/ brown ones are “duds” while the more yellowish planulae are viable (assuming this is consistent across all sun coral colorations).
(For more reading on sun coral reproduction, please see the references below.)
Although the tank wasn’t exactly clean, both sun corals were appreciating the high nutrient levels!
Final Thoughts:
Although I took this photo under actinic lighting, the blue of the sun coral is very similar to the blue of the Oregon Blue Tort Acropora coral. This amazes me as one is photosynthetic while the other is not. In advertising the Oregon Blue Tort, there are plenty of websites that describe the blue as “so vivid it makes you wonder whether it was injected with an artificial blue pigment,” “often considered the bluest Acropora coral you can buy,” “one of the bluest of the blues,” and “so blue it almost looks fake.” I can see why this blue sun coral was also believed to be dyed!
Conclusion:
Over 13 months later, here it is (under actinics again, so the yellowish green polyps are hard to capture on the camera). I lost four polyps, and a few polyps shrank but are now re-growing. The coloration hasn’t changed a bit. The tissue is brilliant blue, the tentacles are yellowish-green, the oral cavity is yellowish-green, and the center of the polyp is blue. Shaded areas turn yellowish-green, but they regain the brilliant blue color once exposed to light again. This makes me believe it’s a structural blue versus a pigment.
Although I lost four polyps, this gives me an opportunity to clean the skeleton to help identify the coral (to be included at a future point). I am convinced that this coral is not dyed, but I have no explanation for its coloration, especially as blue is rare in nature.
So, let’s celebrate this amazing coral with a happy ending. It was fate that on 25 July 2021, the Washington D.C. Area Marine Aquarist Society (WAMAS) hosted a virtual meeting with guest speaker, Matt Wandell, from the Monterey Bay Aquarium. His topic was on “Use and Care of NPS Corals.” Of course, I had to chat with him about this unusual beauty. I am happy to report that we have made arrangements for at least half of the coral to go on display at the Monterey Bay Aquarium in the fall of 2021 (once the weather cools down).
References:
Arthur Jen, 2017 (original post is now deleted). https://www.reef2reef.com/threads/baby-jb-for-trade-southern-california.333673/
BSA Corals, “Blue Emerald Sun Coral!,” 26 June 2020. https://www.reef2reef.com/threads/blue-emerald-sun-coral.733669/
Calfo, Anthony, “The Tragedy of Artificially Dyed Live Corals,” September 2002. http://www.reefkeeping.com/issues/2002-09/ac/feature/index.php
Fatheree, James, “Aquarium Corals: A Look at the Sun Corals,” Advanced Aquarist, 2011. https://reefs.com/magazine/aquarium-corals-a-look-at-the-sun-corals/
Mizrahi, D., Navarrete, S. A., and Flores, A. A. V., “Groups travel further: pelagic metamorphosis and polyp clustering allow higher dispersal potential in sun coral propagules”, Coral Reefs, vol. 33, no. 2, pp. 443–448, 2014. https://core.ac.uk/download/pdf/37522324.pdf
ReefdUp, “Blue Sun Coral Spawning!” 12 December 2020. https://wamas.org/forums/topic/95817-blue-sun-coral-spawning/?_fromLogin=1
Rowlett, Joe, “A Field Guide to Sun Corals,” 2015. https://reefs.com/magazine/field-guide-sun-corals/
Tennyson, “How to tell if a coral is dyed a different color?” 6 July 2020. https://www.reef2reef.com/threads/how-to-tell-if-a-coral-is-dyed-a-different-color.736716/
Tennyson, “HUGE Sun Coral Spawning event,” 9 October 2019. https://www.reef2reef.com/threads/huge-sun-coral-spawning-event.654328/
Wells, CD, Sebens, KP, “Individual marking of soft-bodied subtidal invertebrates in situ – A novel staining technique applied to the giant plumose anemone Metridium farcimen (Tilesius, 1809),” PLOS ONE 12(11): e0188263, 21 November 2017. https://doi.org/10.1371/journal.pone.0188263
Montipora-Eating Nudibranchs and Treatment
Growing…growing…gone. Unfortunately, this is an all-too-common story in the reefkeeping hobby due to a pest aptly named for its favorite meal, the Montipora-Eating Nudibranch (Phestilla subodiosus). Although treatment of Montipora-Eating Nudibranchs is historically challenging, this article covers several available options.
Montipora-Eating Nudibranchs (M.E.N. – just kidding, I won’t use that acronym) are from the phylum Mollusc, class Gastropod, sub-class Opisthobranchia, order Nudibranchia. From there, they are of the suborder Aeolidina, superfamily Fionoidea, family Trinchesiidae, genus Phestilla, species subodiosus. Identification down to the species level was only accomplished as recently as 2021, although the hobby has been actively fighting them since at least the early 2000’s (Wang, et al., 2020).
How to Identify:
These nudibranchs are white in color and range in size from 1-3mm in length. The body is covered in cerata, which may store the nematocysts (stinging cells) of the coral to help protect them from predators. Shown in the picture is a single Montipora-Eating Nudibranch (circled in red) at the edge of a Montipora digitata coral.
The key to detection is to understand the life cycle of these nudibranchs. If there is only one nudibranch, it is capable of reproducing asexually. The adults lay tiny clusters of 3-20 eggs (Borneman, 2007) in crevices along the base of the coral (see image). As the eggs hatch, the juveniles immediately join the adults in eating the coral in a distinct jagged line along the base and edges. Reportedly, these nudibranchs can survive extended periods with no food source.
Back in grad school, I modeled the lifecycle of these nudibranchs and how various treatments worked on them. Unfortunately, the results were never promising once the nudibranchs established themselves in a mature tank. (Note: Sadly, I built this model with an educational license, and my files are “legacy file types” for the software. I have been unable to regain access to my work for either historical purposes or additional research.)
Montipora-Eating Nudibranch Treatment Options:
Treatment for the Montipora-Eating Nudibranchs comes in the forms of prevention, natural predators, chemical destruction, and manual removal.
Treatment Method 1: Prevention of Montipora-Eating Nudibranchs:
Prevention lies in purchasing from a reputable dealer or hobbyist first and foremost. Chances are, if the person has had them before, that person will have them again. Next, the responsibility falls to the new owner of the coral in proper quarantine procedures (at least two months is recommended). Examination with a magnifying glass is recommended to help identify and remove eggs. However, the nudibranchs might have laid eggs in a hidden crevice. Since the eggs are laid at the base of the coral or on dead coral skeleton, some hobbyists chose to remove the coral from the base rock. Both remaining coral on the rock and base rock then dies. This is to help ensure that the eggs are not transferred.
Treatment Method 2: Controls for Montipora-Eating Nudibranchs
Some hobbyists have found certain natural predators will help control the nudibranchs. Unfortunately there are no known complete eradication measures available naturally. The wrasses generally known to eat most invertebrates will most likely also feed on the nudibranchs. It is important to keep in mind that as far as natural predators go, what works for one hobbyist may not work for another hobbyist, especially with wild creatures.
Treatment Method 3: Coral Dips for Montipora Eating Nudibranchs
The last method of eradication is by chemical means. Coral dips often help stun the pests so they can be blown off gently with a pipette or powerhead. These adult Montipora-Eating Nudibranchs (as pictured) were in a dip much longer than the coral colony could survive. Unfortunately, that is typically the case; however, there are a few less lethal options.
Montipora-Eating Nudibranch Treatment: Iodine
Many hobbyists have found that an hour-long dip in iodine will kill most adult nudibranchs. However, this may cause undue stress to the host coral, especially if flow, pH, and temperature are not ideal in the dip.
Montipora-Eating Nudibranch Treatment: Levamisole
Levamisole (a pig dewormer available at farm supply stores), has been used in a similar manner to the iodine dip, but with a higher fatality rate to the host coral. Levamisole paralyzes the nudibranchs so they can be removed more easily from the coral.
Montipora-Eating Nudibranch Treatment: Potassium Permanganate
Another chemical method is potassium permanganate. The dip (50mg/l for 30-90 minutes – Borneman, 2007) has been shown to effectively destroy both the adults and the egg masses with moderate impact to the coral. After eradication on the corals, the main tank should be kept free of Montipora for approximately two months.
Like all chemicals, caution should be exercised when handling potassium permanganate. Using an MSDS (Material Safety Data Sheet) to learn about the hazards of use and what protective gear is highly recommended. An MSDS for potassium permanganate suggests the use of eye protection, gloves, and respiratory protection as potassium permanganate can cause serious burns to the skin, nose, and throat.
Montipora-Eating Nudibranch Treatment: Series of Dips + Manual Removal
Rather than trying to kill both adult nudibranchs and eggs, I attempt to just break the lifecycle by removing the adults so that they cannot reproduce. When I get any frags of Montipora, I perform my usual coral intake process. I perform an inspection, remove the plug/base/dead skeleton, perform dips with manual pest removal, and attach a new base. Then I place the coral into my quarantine tank, but it stays for an extended period (60 days rather than the typical 30 days).
Since a succession of coral dips work for AEFW and red bugs, I also dip all Montiporas daily the first week to aid in manual removal. After that, I dip every other day the second week, every third day the third week, and then once a week for weeks four through six. I have not lost a single coral with this method, and it saves me the hassle of treating with different medications for different parasites at different times.
Various dips may work to stun the nudibranchs, and examples include CoralRx, Revive, and Tropic Marin Pro Coral Cure. Again, here the goal is to simply aid manual removal by stunning the adults. If the adults are removed, the eggs will continue to hatch in the presence of food. Adults are then easily removed before they can reach maturation. In the lifecycle model mentioned above, this was the most successful approach when implemented correctly.
Montipora-Eating Nudibranch Treatment: Display Tank + Dips
The worst challenge is a display tank with mature colonies infested with nudibranchs. My best advice is to consider the coral dip series described above (although large colonies may require buckets for treatments). Additionally, the hobbyist would have to kill any bases left attached to the rock along with unremovable encrusted Montipora colonies. Another option, if the colonies are smaller, may be for the hobbyist to remove all Montipora from the display tank. The hobbyist would then place all removed Montipora into a quarantine tank for treatment, minus a sacrificial frag that stays in the display. The hobbyist must then frequently replace that sacrificial frag so as to draw out the nudibranchs from the rockwork.
Montipora-Eating Nudibranch Treatment: Other Dip Options
Hobbyists have used garlic and freshwater dips with nudibranch mortality but they also often report high coral mortality.
Conclusion:
Although there are still no full-tank chemical treatments, eradicating Montipora Eating Nudibranch is possible with patience and proper husbandry. Using a combination of dips and manual removal seems to be the most effective option available at this time.
Red Bug Parasites (Tegastes acroporanus)
Somehow back in the day, after over five years of having SPS-dominant aquariums, I got my first case of red bug parasites. A local aquarist was tearing down his system and gave me a few corals that weren’t doing well. Turns out, red bugs AND Acropora-eating flatworms (AEFW) infested the corals. Fortunately, I caught the red bugs while the Acropora was still in my quarantine tank.
Red bug parasites (Tegastes acroporanus) are extremely tiny copepods (as seen in the red circle in the photo) that solely infest Acropora corals (not Montipora, Pocillopora, etc.) They come into the hobby on wild or maricultured colonies (corals purposefully grown and harvested in controlled ocean environments), and hobbyist selling/trading of aquacultured corals further spreads them.
A good indicator of a red bug infection is a loss of Acropora tip coloration and browning that cannot be attributed to other factors. Red bugs caused the coral shown to lose nearly all coloration, while AEFW caused the white patches.
In the early 2000’s (now I’m dating myself), there was a general consensus in the reefkeeping hobby that red bugs only affected smooth-skinned Acroporas. Hobbyists assumed they did not bother corals like Milleporas or “the green slimer” due to the extra mucus they produce. Although red bugs do tend to prefer certain Acropora (especially Acropora valida, the “tricolor”), most every Acropora species is susceptible. Hobbyists also commonly believed that red bugs laid eggs and had a larval stage. This led to a consensus that three treatments were needed in order to eliminate the red bugs entirely. Although research eventually determined that red bugs are live bearers, I still recommend three treatments just as a safety precaution.
How to Identify:
On Acropora corals that have “browned out” due to red bug presence, the red bugs are rather distinguishable. They are mostly yellow (almost a gold color) with a bright red dot. They are also extremely tiny at around 0.5mm (think a little bigger than the size of the period at the end of this sentence). Unfortunately, they are not easy to see on healthy corals, and they have an ability to hide within the coral polyp itself without being consumed. In order to determine the presence of red bugs, I use a magnifying glass to look for them before placement into my quarantine system. After that, I observe the coral daily for a week. I also take a photo of each coral and zoom in to look for anything I’ve missed.
How to Treat Red Bug Parasites:
Unlike other serious pests (e.g., Montipora-eating Nudibranchs), there is a suitable in-tank treatment for red bug parasites.
Method 1: Control
Some hobbyists believe that blowing the corals off with a powerhead a few times for a couple weeks will rid the Acropora of red bugs since they may starve before finding their way back to the coral. Other hobbyists rely on natural predation (e.g., pipe fish, file fish, gobies, wrasses), but to my knowledge, no predators have a 100% success rate. I advise against natural control methods for aquarists who plan to sell/trade their corals.
Method 2: Coral Dips
Coral dips often help stun the pests so they can be blown off gently with a pipette or powerhead. Since red bug parasites are live bearers, this method can work as long as all the red bugs are removed. Since a succession of coral dips work for AEFW and red bugs, I dip all Acroporas daily the first week. After that, I dip every other day the second week, every third day the third week, and then once a week for weeks four through six. I have not lost a single coral with this method, and it saves me the hassle of treating with different medications for different parasites at different times.
I should note that I’ve never found a pest after week two, but I still go the full six weeks since the lifecycle of AEFW is quite long (and I always treat for both AEFW and redbugs). Also, some dips can be extremely damaging to smooth-skinned corals, so please use caution if you plan to treat one of these.
Coral Dip Treatment Protocol for Red Bug Parasites:
Follow dip manufacturer instructions.
Method 3: Prescription Flea Medications
Before the manufacturer discontinued it, hobbyists considered Interceptor (milbemycin oxime), a prescription canine flea medication, as the golden standard for red bug parasite treatment. However, as it killed all shrimp, crabs, and other crustaceans, aquarists had to remove them before dosing (the pod population recovered with time). This die-off would also spike ammonia levels, so reefkeepers had to be prepared for a water change and to run carbon. Many people had difficulty obtaining Interceptor from their veterinarians since treating for red bugs is an off-label use. However, the Veterinary Information Network (VIN) published information on red bug parasites for reference.
Once Interceptor was no longer available, reefkeepers began searching for a replacement. Currently, Milbemax is the most often recommended replacement. It is a prescription combination of milbemycin and praziquantel. (The latter medication is commonly used for fluke treatment in reef tanks (e.g., PraziPro); however, it is a much lower dose than would be effective for common marine flukes).
Providing proof of infection to a veterinarian and asking him/her to look up red bugs on the VIN should be enough to receive the medication (or at least it was for us.) Some veterinarians have recommended different medications with different treatment protocol (e.g., MilbeMite Otic), so those individual cases are not discussed here. Unfortunately, many aquarists are finding some red bugs have a resistance to milbemycin. If you try it with no success, work with your veterinarian to find an alternate medication or try one of the other methods listed here.
Interceptor Treatment General Protocol for Red Bug Parasites:
(This is provided for historical purposes only since Interceptor is no longer available, as it may be relevant to other treatments.) Dustin Dorton of ORA determined 0.025 grams of Interceptor are needed per 10 gallons of water. After that, the protocol is the same as Milbemax, described below.
Milbemax Treatment General Protocol for Red Bug Parasites:
(Obtain actual procedures from a veterinarian, as this is for general reference only.) The recommended dosage is 23mg per 400g tank water, which works out to 0.0575mg per gallon tank water (or 5.75mg per 100 gallons if that is an easier reference point). Similar to Interceptor medication, pulverize and weigh the medication out as accurately as possible. Estimate the true volume of water as close as possible (subtracting live rock, sand, etc.)
Remove all crabs, shrimp, and other crustaceans from the aquarium before dosing. Turn down the skimmer so that water runs through it, but the skimmer doesn’t actually skim. Remove GFO and activated carbon. Add an airstone (or multiple, depending on the size of the tank) bubbling gently to help maintain the pH. Otherwise, allow everything else to run so that the chemical can spread throughout the aquarium. Dose the correct amount and let the system run for six hours.
After six hours, the red bugs should be gone, but if they are not, slightly adjust the dosage and try again in a few days. Perform a water change (~15-25%), turn the skimmer back up, and bring the GFO and carbon back online. As mentioned above, although red bugs are live bearers, three treatments a few days apart is still ideal. Once the red bugs are completely eradicated and the medication is removed, reintroduce the crabs, shrimp, and other crustaceans to the aquarium.
Method 4: Bayer Advanced Insecticide
WARNING: Per Bayer Advanced Insecticide’s labeling, “It is a violation of Federal law to use this product in a manner inconsistent with its labeling.”
Yes, this is the stuff you may have in your garage or at your local hardware store. Some aquarists use it as a dip only, not as an in-tank treatment. This website does not cover Bayer use as a coral dip, due to the warning above.
Method 5: Temperature Adjustment
With crazy heat in the summers, lowering the tank temperature may not be feasible. But for tanks located in basements during the winter, this may be the least expensive option. In fact, this method worked for me…accidentally! Right after I received the infested Acropora mentioned above, we lost power for about six hours. My quarantine tank got down to about 65 degrees (thanks to being in Utah in the middle of winter). After the power came back on, there were no signs of red bugs. I’m not the first to document this. I still continued treating for them just in case, but I never saw another one. This seems to work, but we still need scientific studies to back it up.
Experimental Temperature Treatment Protocol for Red Bug Parasites:
Slowly lower temperature to 65-66 degrees. If tank inhabitants start to show signs of stress, start bringing the temperature back up. Leave the temperature at 65-66 degrees for 4-6 hours. Note: this is experimental, and there is not enough data to determine if this treatment works 100% or is safe. Use with caution!
Conclusion:
Thanks to all the courageous mad scientist reefkeepers out there, there are several proven treatments for red bugs, including an in-tank treatment. The Acropora shown was saved from both red bugs and AEFW! Hopefully, with the increasing ease of treatment protocols, red bugs will be a thing of the past. Best of luck in your treatment!
Dying Lithophyllon Coral Rehabilitation
A local fish store (LFS) called me to ask if I wanted to try to save some of their corals. An employee had overdosed their reef tanks with a chemical used to treat flatworms (I was not told what chemical it was). When I arrived, honestly, I was shocked at how bad many of the corals were. I don’t know if these corals were dying from an overdose of some unknown chemical or if the flatworm toxins were killing them. Either way, something was poisoning these corals to death. I picked up two “chalice” corals, amongst others. These dying Lithophyllon coral mouths were gaping open, the tissue was falling off, and what tissue did remain was barely hanging on. Rehabilitation was going to be dicey. (Photo of Litho#1 taken 11 May 2013)
Dying Lithophyllon Coral Inspection and Dipping:
When I brought them home, I temperature acclimated them to my quarantine tank and prepared the dips. Like most of my rescues, I dipped the corals for pests. (Photo of Litho#2 taken 11 May 2013)
*NOTE: This post tracks the corals as “Litho#1” and “Litho#2” to show individual progress.
Coral Quarantine:
After the dips, I placed the two dying Lithophyllon corals into my quarantine tank in a low light, low flow area. I made sure that no crabs would bother the coral, especially with all the decaying tissue. Since the corals were poisoned, I made sure the quarantine tank had new carbon to help absorb anything that the corals might leach. Both corals did start to quickly recede, so I tried to stop it by smearing a bead of superglue gel along the receding edge (white lumpy band in the photos below). Rehabilitation was definitely already proving rough. (Photo of Litho#1 taken 11 May 2013)
The corals also refused to eat until about a month after I received them. I started them on fish food pellets (shown here and above) since they are quite processed. In my experience, sick corals seem to digest fish pellets better than whole foods (like mysis, etc.) This helps them recover faster. (Photo of Litho#2 taken 9 June 2013)
Coral Grow-Out:
After stopping the recession with superglue and maintaining regular feedings, the corals started to heal quickly. And how great they started looking! After 30 days in quarantine, I inspected, re-dipped, and placed them into the grow-out tank. (Photo of Litho#2 taken 4 August 2013).
Regular feedings likely helped the coral recover. (Photo of Litho#1 taken 12 October 2013)
Here is Litho#2 on 3 November 2013.
A month later (8 December 2013), Litho#2 was growing down its skeleton.
Fully Rehabilitated Lithophyllon Coral:
Nearly a year later, both previously dying Lithophyllon corals were growing rapidly. This Litho#2 is shown on 1 May 2014, when it was fully rehabilitated. Below is a close-up shot showing the corals’ beautiful patterns.
Conclusion:
I had very little hope for these dying Lithophyllon corals due to their poisoned state. However, the fast reaction of the LFS along with continued care over a year fully rehabilitated these two corals. Shown below are the before/after photos of the dying Lithophyllon coral rehabilitation. One coral was nearly immediately rehomed, and the other coral was rehomed a couple years later.
Dying Cyphastrea Coral Rehabilitation
Cyphastrea coral are usually very hardy and prefer low light (and sometimes do best partially shaded) but can be acclimated to higher-light areas. They also adapt well to a variety of flow conditions. I received this dying Cyphastrea coral from a reefkeeper who was getting out of the hobby. Good husband would likely be key to this coral’s rehabilitation. (Photo taken 22 May 2012)
Unfortunately, most corals in the tank were suffering from poor water quality, and this Cyphastrea was no exception. The reefkeeper did the responsible thing by recognizing that the corals needed proper care and attention.
Note how some of the polyps still have tissue while there is no tissue between the polyps. This is a good indication of poor water quality (in addition to actually speaking with the previous owner). Regardless of the damage root cause, I still dipped and checked it for pests.
Coral Quarantine:
As mentioned, the coral was declining due to poor water quality. After the initial dipping process, I placed the coral into my quarantine tank for observation and acclimation. Here, the coral was fairly low in the tank to allow it to adjust to the lighting. Notice that by less than a month later (15 June 2012), the coral was already regrowing!
Just another 15 days later (30 June 2012) and slightly over a month total time with me, and the coral was gaining coloration and growing rapidly! Given the proper light and water conditions, this coral grows extremely
fast.
Fully Rehabilitated Coral:
This previously dying Cyphastrea coral was completely rehabilitated in less than three months (pictured at 14 August 2012). The coral had completely covered the old skeleton and had encrusted onto the base. In fact, it started to grow so quickly, that I had to frag it several times. Coral reproduction via fragmentation reduces hobbyist impact on the oceans.
Troubleshooting Dying Cyphastrea Corals
As mentioned above, if the Cyphastrea coral has started to lose tissue between the polyps, then there is probably a water quality issue. If the coral is bleaching, then it’s likely in too high of light (or might’ve been in absolutely no light.) If the coloration isn’t great (too brown), then it may need more light or less nutrients. Or, if there’s recession from one side to another, then it may be a pest, bacterial/fungal infection, or a sudden water quality problem.
