Breeding Journal, Species: Pseudochromis fridmani (Orchid Dottyback)

General
Species: Pseudochromis fridmani (Orchid Dottyback)                  
Social Structure:  Male/female pair
Size of Individuals:  Female ~2 inches, male ~3 inches
Age of Individuals:  Unknown
Date added to Tank:  3/8/2013

Broodstock Tank Details
Size of Tank:  20g High on a 200g+ system
Substrate Details:  None
Filtration Details:  Skimmer, bioballs, live rock
Water Changes:  5-10% every two weeks
Water Temperature:  80F
Lighting:  LED light puck
Lighting Cycle:  6AM to 8PM
Other Tank Inhabitants:  A little crab, species unknown, a couple hermit crabs

Broodstock Feeding Details
Food Types:  Frozen foods including krill, squid, mysis; live blackworms (the female will not eat them)
Feeding Schedule:  2-5x per day

Spawning Details
Date of First Spawn:  4/21/2013
Spawn Time of Day:  ~6PM
Dates of Consecutive Spawns: 
5/1/2013: Eaten?
5/7/2013: Spawned before 6PM, egg ball removed immediately for artificial incubation since the tank has copper in it. ~50 larvae hatched 
5/12/2013: Spawned between 5 and 7 PM, egg mass broke apart before hatch 
5/20/2013: Not collected
5/26/2013: Egg ball taken ~12 hours post spawn, sterilized and tumbled
7/15/2013: Egg ball immediately taken and tumbled in inverted 2L soda bottle
7/22/2013: Egg ball immediately taken and tumbled in inverted 2L soda bottle
Courtship Details:  None noted, but likely typical dottyback spawning behavior involving the male dancing and attempting to lead the female back to his chosen spawning site.     
Egg Size:  ~1mm
Egg Color:  Clear/white-ish
Egg Count:  Not counted

Hatch Details
Hatch Date:  5/10/2013 from the 5/7/2013 spawn
Hatch Time of Day:  all day (artificial egg incubation)
# Days after Spawn:  3 (hatch is on the 4th night)   
Larvae Description:    ~3mm, transparent, no yolk sac
Consecutive Hatch Dates:    7-18-2013 (7-15-2013 Spawn), 10-19-2013,10-25-2013, 11-17-2013           


Larval Tank Details
Temperature:  ~80F 
Size of Larval Tank:  17 gallon BRT    
Substrate Details:  none
Other Tank Decor:  airstone 
Filtration Details:  live phyto
Lighting:  Fluorescent strip light  
Lighting Cycle:  24/7
Water Changes:  ~25% on Day 5

Larval Feeding Details
Food Types:  Rotifers constantly, nhbbs added starting around day 8 or 9; Apocyclops, Tisbe, and Euterpina copepods added occasionally, enriched bbs added after day 17  
Feeding Schedule:  constant

Metamorphosis/Settlement
Date of Settlement Start:  11-24-2013
Days after Hatch:  36
Date of Settlement End:  11-26-2013
Description of Fry:  Miniature adults, about a centimeter long, bright purple with a black line by the eye

Grow-Out Tank Details
Temperature:  78-80F
Size of Grow-Out Tank:  13 gallons on a ~115 gallon system
Substrate Details:  None
Other Tank Decor:  Few PVC pipes, yarn mops
Filtration Details:  Live rock, sponge filters   
Lighting:  Fluorescent strip light
Lighting Cycle:  6AM to 9PM
Water Changes:  Occasional and random
Size at Transfer:  ~2 cm
Age at Transfer:  ~ 1 month post settlement (Transferred 12-29-2013)

Grow-Out Feeding Details
Food Types:  Enriched bbs, golden pearls, crushed flake, Spectrum grow pellets, frozen, enriched artemia 
Feeding Schedule:  Two or more times a day

Additional Information
 
(No Pictures or Videos in the Section Please)
Miscellaneous Information: 
The male is wild caught, the female is ORA captive bred.
 
The first settlement I got was from the October 19th hatch.  The first settled juvenile was seen November 24th and the last larva in the tub was November 26th.  I ended up finding three settled when I pulled the sponge filter and heater out.


You will be required to provide photographic or video evidence in this thread of each event submitted for the MBI Program.
If your thread does not contain these photos the MBI Committee will not be able to approve your reports. PHOTOS AND VIDEO S MUST BE PLACED IN ADDITIONAL POSTS, NEVER IN THE FIRST POST IN A JOURNAL.

Here is a quick picture of the parents.  They are actually pretty hard to photograph because the LED light puck above their tank isn't very bright, so then I have to use the flash, which scares them a bit.  I don't want to make them too nervous now!

 
Here is the egg ball in a 1" PVC tube that is about 7" long

I was surprised at how big it looked considering the female just seems so tiny
 
I was a bit surprised with this spawn. The female had been looking a little gravid, but I was thinking it was more due to her starting to eat way more aggressively lately than she had been.  I was expecting my neon dottys to spawn this weekend (it seems they had a missed connection instead) so this is a fun surprise.  A friend also gave me some Hippocampus comes seahorse fry to try my hand with on Friday, so I may have my hands full in this egg ball hatches!

The egg ball is still there as of this morning (4-25-13)
 
I am shocked (and pleased!) that he did not eat it so far.  Fingers crossed he keeps taking care of it today since they will hopefully hatch tonight.  The egg ball seems a bit smaller than it was originally, but not too much smaller.  It is hard to really say because I have been trying not to spook him too much with the flashlight.
 
Also, the egg tube has been moved twice since they spawned.  The first time, I moved it and propped it up during a water change that I did immediately after the spawn (the tube is long and the first time I looked I didn't see eggs...so I went ahead with the scheduled water change and their tank was due to be cleaned; good thing I checked again!).  The second time the male must have knocked it down or something because in the morning it was no longer propped up on top of another PVC section, but had rolled several inches away.  Luckily, neither of these tube rearrangements have bothered him much.

In the future be very careful about moving the pipe, it may cause the eggs to be eaten. I never messed with the tanks while eggs were around. If I did the eggs were usually eaten.

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

I didn't intentionally or knowingly do it. I didn't think they had spawned when I cleaned the tank, and the second time wasn't anything I did or was involved with. I just wanted to make note of it was all

I know, I was just saying be careful in the future. Once they get on a schedule you'll be able to tell when they will spawn. Mine were on a nice 7 day cycle so I could clean their tank a day, or so after the eggs hatched.  

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

My male used to be really sensitive, it was like you look at his egg ball through the glass and he would eat it.  Now I can actually remove the egg ball to inspect it, and put it back and he won't eat it.  That's pretty unusual in my limited experience.

Don't let fear and common sense stop you! =]

Of all the problems I've had with my P. tapeinosoma and P. cyanotaneia, the male eating the eggs has not been one of them.  I, too, have removed the egg ball to sample a couple under the microscope, and returned it, and even lost track of the number of days, and tried to hatch them in the tumbler one day early once, then returned them to my P. tapeinosoma male, and he continued to properly care for them.  Every time the P. cyanotaneia spawn, the female exits the cave prematurely, and drags the egg ball with her, and I have to find it and give it to the male at the entrance to his cave, where he takes it and puts it in the back of the cave where it belongs.  I also examine them with a flashlight regularly.  None of this has ever phased my males.

Well, I have no larvae and no egg ball.  I was working late last night so I had put the snagger in yesterday.  Either they hatched during the day (a friend didn't flip the snagger "on" until about 6PM), he ate the egg ball on the day of the hatch, or they hatched and the snagger missed them (when I checked at about 10:30 last night the snagger suction cups had slipped and there was an escape route out of the snagger the larvae could have taken).  
 
Regardless, no larvae
 
Hopefully they will spawn again on Sunday.

Another spawn last night on 5/1/2013, 10 days since the last spawn.  I think part of that is due to the larval snagger that was in the tank for a day.  The male did not seem to like it at all and hid for a couple days even after it was removed.  Egg ball looked to be about the same size as the last one.

The egg ball was much smaller as of yesterday, perhaps half the size as it was originally, if not smaller. Hopefully I will still have some eggs by the time the hatch is due!

Eggs are gone

Another spawn today, which was surprising for two reasons. One, it seems a bit early, and two, there is copper in the tank. Because of the second reason, I actually took the eggs out right away and put it in a flask to artificially incubate (or attempt to, at least).
The egg ball:


The flask:


Not sure if the eggs are fertile or not. We shall see!

Today I disinfected the eggs since I had not done it yesterday (all I had done was put them in new saltwater). I used 5mL hydrogen peroxide in 1L of water for 15 minutes while keeping the egg ball gently tumbling around. Then I scooped the ball out with a very fine net and put it into fresh saltwater again in the flask. I have the flask sort if protected from light by wrapping it with gray WalMart bags (looks ridiculous but it works. Sort of).

There were a few free floating eggs in the flask but they looked white so I am guessing they aren't any good anyway and I did not scoop them out with the egg ball.

I wonder if I should sterilize the egg ball again tomorrow if it makes it until then?

The egg ball started to really break apart during the night.  Yesterday there were definitely more free floating eggs and it was clear that only about 1/3 of the egg ball was developing.  During the first night of tumbling, the egg ball had settled in the flask and I wonder if the non-developed eggs are the ones that settled on the bottom and didn't get as much water movement and died.  Or, perhaps the male didn't fertilize them to begin with.  I didn't check with the microscope when I first pulled them.  There was one free floating larva, but it did not seem alive. Considering the way this egg ball is rapidly breaking apart, I don't anticipate getting much out of it.  I will clean out the flask today again just in case though.
 
The female is looking gravid again, so hopefully I will be able to tweak the egg tumbling set-up and getting better results with the next egg ball.

Well, some of the few remaing eggs hatched today/tonight!

See the eyeballs??

And under the scope:


There are probably only twenty or so. I took them and put them in a few gallons of water in a 10 gallon tank, added some T-iso, rotifers, and apocyclops. I don't have much hope, but 15-20 larvae are better than none

Yay!  It's a start.  Congratulations!

Another spawn already today (5-12-2013). I accidentally caught them in the act, and by the time I went back down and could tell the female was back in her usual tube, the male had eaten the eggs. I think I made him cranky by stealing the eggs last time and then interrupting them this time :/

Fake out. I found some eggs A small spawn, but I'll take it. I put it in my new tumbler based on fin farm's tumbler. I put the tumbler in a 2L water pitcher with aged new ASW and two drops of methylene blue. I may dilute that out a bit yet tonight, not sure. The tumbler is pretty cool because the eggs just sort of hover in place. I put black electrical tape around the water pitcher to keep the eggs dark and clamped it in to a broodstock tank.

Here's a pic of the tumbler:


There are a few larvae alive still from the hatch that just happened on Friday cruising around. I have left the lights on 24/7 over them. They seem so small!

Should I be concerned that the time between spawns went from 10 days to 6 days to 5 days?  5 days between spawns seems super short... I mean, its nice to have another chance so soon, but it is also a bit worrisome.  Could this be an effect of the copper?

Mine stuck to a 7 day cycle most of the time. As long as yours are spawning I wouldn't worry about it.

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

The last egg mass got all fungus-y by the day of the hatch. I will try the same tumbling method again, but keep the flow higher in the beginning and sterilize the eggs with hydrogen peroxide the first two days (maybe the third too) instead of leaving methylene blue in the entire time.

No spawn yesterday, but I am hoping this weekend I will get another egg ball to play with.

The last larva from the hatch on 5-10 disappeared yesterday.

There was another spawn last night, but it was a super small egg ball.  I may have had better luck tumbling a small egg ball because there are just fewer eggs to keep oxygenated in the center, but I decided to just let the male keep this one for practice.  This spawn was 7 days since the last one.  

Another spawn on Sunday, May 26th. I didn't remove it immediately (I forgot) but I took the egg ball this morning so the male had it for about the first 12 hours, and during that time it was exposed to copper. I am not sure how detrimental that will be. I sterilized using hydrogen peroxide (2.5mL in 500mL of ASW) for 15 minutes and then put it in the tumbler set-up. The egg ball is small-ish and stays at the top of the tumbler with very little air, whereas the last one stayed toward the bottom with high air flow pulling water through.

Well, yesterday it became clear that the eggs were bad (they were white).  So yet another tumbling failure.  Not sure if the problem this time was the actual tumbling itself, or if it was because the eggs were left in the copper tank overnight.
 
Luckily, the copper will start being removed from the broodstock system next week or the one after, so hopefully I won't need to steal the eggs from him in the future.

Another spawn today. I removed it very shortly after they spawned and the male put up a bit of a fight for them. This time I am doing something a little different. I put the tumbler in the 3 gallon (or so) fish bowl/orb with a heater and a bag of cuprisorb. I put two drops of methylene blue in also. We shall see if the additional water volume and the cuprisorb help.

They have been spawning reliably every 7 days, with the exception of one week where it was 8 days apart.  So now I think they usually spawn on Mondays instead of Sundays.  Monday (about 6 PM or so) I pulled an egg ball, but it is possible that they spawned on Sunday.  I rinsed in fresh ASW and sterilized using hydrogen peroxide, then washed again and then put them in an inverted 2L soda bottle (cut in half, with the bottom being used as a base).  I drilled a hole in the cap just large enough for airline tubing to fit through and pushed it about 1/4 inch into the "tumbler."  The egg ball stayed in pretty great shape and tumbled better than I expected.  The morning of the 18th I woke up to a completely broken apart egg ball but I just left the eggs tumbling.  I checked the eggs again around 7PM on the 18th and noticed several had hatched.  By 10PM the majority of the good eggs had hatch and the larvae were put into a BRT with about 7 gallons of water in it (17 gallon capacity), an airstone and a strip light over it (left on 24/7 at this point).  The tub had been running for about a week and has a great population of rotifers, and several different types of pods cruising around in it (just tisbe, psuedocyclops, tigriopus, and maybe one other that are cultured as a polyglot).  The water was tinted with nanno and chaetoceros.
 
This morning there were plenty of larvae cruising around and in the few minutes I took to check on them, I saw several striking at food  I would say there are at least 100, but it's sort of a wild guess.

Congrats,Shannon!
I am letting the father take care of incubation,but eggs vanish somewhere in the 1-3 days range.Wonder if I should start pulling/tumbling them

 
If the eggs are disappearing anyway, then I think it's worth a shot tumbling them.  Even when the egg mass broke apart I could tell there were still some good eggs and they still hatched.  I ended up with over a hundred larvae to tinker with, which is far far better than none  I have tried three different tumblers so far.  The first "tumbler" was just a flask with an airline in it.  It worked, sort of, but less than 50 hatched.  The next was more of an airlift type tumbler made from parts of a sponge filter and I think that had the most potential, but I am pretty sure my plastic tumbler parts got contaminated with copper or something because the eggs always died a couple days in.  This last attempt with the 2L bottle worked better than I expected and it's what I will likely do with the egg ball I should be getting tonight.  I will likely try to build a new airlift type tumbler out of PVC and clear tubing sometime soon-ish, but not a priority at this point.
 
The larvae are in Day 4 right now, and still seem to be doing well.  I can't tell if there has been a lot of mortality since they are in the BRT and its difficult to see stuff on the bottom even with a flashlight, but it doesn't seem like the numbers have dropped significantly.  Can anyone tell me when the starvation point is for these guys?  I think I will be on edge until I know that that first critical point is passed.  Or is every day a critical day with these?  I see them striking at food, but they don't seem to have that peanut shape from the top that clown larvae get.  Is that bad? I suppose I could pull some out and see from the side if they have silver bellies.
 
I have re-read Moe's Orchid Journal book and on some runs he would feed nhbbs as early as a couple days, in small amounts.  Has anyone tried that?

Another egg ball last night, spawned before 7PM. I pulled it, rinsed it, sterilized using 2.4mL of hydrogen peroxide in 475 mL of ASW, rinsed again, and put it in the inverted 2L bottle tumbler.  This morning the eggs were on the bottom and the air seemed to have slowed down so I turned it back up and got the egg ball moving again.  As of this morning it still looked like most of it was alright.
 
The larvae from the first hatch are on day 5 now and *seem* to be doing well and eating. 

Yesterday (Day 5) I did a water change/bottom siphon.  It was actually pretty easy to avoid sucking up any larvae because none of them cruise around the bottom.  I pulled out about two gallons (I estimate there is about 7 or 8 gallons in there) and only sucked up one larva.  I did, however, remove a TON of copepods which is sad.  So I ended up adding some more pods from the polyglot culture to hopefully make up for it.  It would be great if the pod population was good enough to keep them going without having to add a lot (or any) BBS.  I added 3 gallons of freshly mixed ASW back in via  airline tubing (no dripping...just draining quickly).  
 
I saved the water that I removed so I can harvest the pods and rots from it and use it to seed a new BRT for the next hatch that I hope to get.  The tumbling eggs still look good and I am hoping to see some eyes develop sometime today. If the eggs go bad, then I will try to get mainly just the pods out of the waste bucket and either add them back to the BRT with the dotty larvae or use them for the clingfish tub.
 

Hard to really see, but here are some of the larvae cruising around in their bucket:

The egg ball looks shot.  No eyes ever developed and nearly all the eggs look white-ish, so I tossed them. 
 
The larvae are on Day 7 now.  They got a 5 gallon or so water change (of 7 or 8 gallons).  I siphoned the bottom and then put the siphon in a 400uM sieve so I would pull out only rotifers and no larvae.  I added back in ~5 gallons and also used chaetoceros to tint the water again.  They seem active and strong and I would say that there are still maybe 100.  My original estimation of the number that hatched was definitely way off.  There were probably 200 or so, maybe more.

Shannon,this is a very interesting subject as many breeders are having problems hatching dotty eggs and it is not clear if we must blame egg quality,parent eating them or poor artificial incubation technique.
I have started working on that,but dont mean to HJ your thread:
http://www.marinebreeder.org/forums/viewtopic.php?f=211&t=11853

It's not a hijack at all, since I always have to artificially incubate these eggs...the more information the better!
 
Are you sterilizing the egg ball before you start tumbling?  This last egg ball (from Monday evening) I did not sterilize (it was late when I pulled it and I was lazy. Tsk tsk to me) and it fell apart more quickly than previous attempts when I did sterilize.  So far, I prefer the hydrogen peroxide short term bath over the methylene blue during incubation/tumbling.
 
I plan to make a new (definitely never exposed to copper!) tumbler out of a sponge filter again to try that method next week with the new egg ball. 

I had no problem getting eggs from my original pair to hatch. The male tended to them and I pulled and hatched the eggs when they were ready. Getting them to settlement was more difficult.

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

Tal,some breeders brood their eggs until hatching,while others don´t.Perhaps a matter of luck?

I am artificially incubating because there is copper in the system, otherwise I would be happy to let the male keep them until hatch.  I also have another pair of dottys that spawn occasionally, but the male eats the eggs within 15 minutes of spawning.  So, if I get lucky enough to catch them in the act again, I will steal those eggs and artificially incubate them too.
 
A brief update...the pair spawned yesterday.  The female was HUGE during the morning feedings, and by the last feeding of the day was slim so I checked the male's den, and sure enough, he had a nice size egg ball in there.  I stole it, sterilized for 15 minutes with hydrogen peroxide and put it in the inverted 2L tumbler.  I had planned to make a new tumbler, but the spawn came early and I wasn't prepared, so the egg ball went into the old tumbler.  I turned the air up a bit higher than I normally would to make sure the eggs wouldn't settle on the first night (like I've had problems with before) and luckily this morning they were still tumbling around and intact.  Fingers crossed that they will make it to Wednesday for a hatch.
 
The larvae from July 18th are still going, although the numbers have decreased down to 20 or so.  They are on  day 18 now. The temperature in my house has been lower than I would like (around 74 for much of this run) and the BRT has no heater, so I am sure development has been slower.  They seem to be taking nhbbs and most are quite active, so I am hoping they continue to do well.  I saw one chomp down a copepod, so hopefully they are also eating lots of those up (there are tons of tisbe in there...the bottom is just crawling with them).

Shannon,following your advice,I began stealing/tumbling the egg balls,and indeed eggs were at least partly fertile.But for some reason they die at some time short before hatching.See pics in my thread linked above.
If this is a bacterial issue,there are two ways to explore;keeping the eggs in an antibacterial treated medium or else give them short baths.As I am tumbling with a running system flow,the 2nd choice is the only possible.

 
I would at least give them a short (15 minute or so) hydrogen peroxide bath before you start tumbling.  And if you are tumbling using system water, maybe pull the eggs for a short bath every day as long as the egg ball is intact? I always plan to do this, but then forget to, and once the egg ball busts up there isn't much that can really be done.   Would it be possible for you to tumble in a separate container?  I'm not sure if that would help or not.  Have you looked at this thread http://www.mbisite.org/Forums/tm.aspx?m=82843 particularly fin farm's tumbler set up? 
 
I've looked at your thread, but it won't allow me to see the attached images and I can't remember my log-in info.  I sent a password request but didn't get it yet.  Once I do get it, I will look at the images