Effect of boric acid on the nuclear division and germination of <i>Saprolegnia</i> spores.

<p>Confocal laser scanning microscopy images of <i>Saprolegnia</i> spores stained with the nucleic acid dye DAPI. a1–a4) Spore germination in non-treated water control group. Note the movement of the nucleus towards the newly developing germ tube following 2 and 4 h incubation (a1 and a2). Development of multinuclear hyphae indicating growth and viability is shown in image a3 and a4. b1–b4) Gradual reduction of fluorescence intensity of <i>Saprolegnia</i> spores treated with boric acid following 2, 4, 6 and 8 h of incubation, b1, b2, b3 and b4 respectively. No nuclear division was observed in the treated group.</p

Viability of treated <i>Saprolegnia</i> spores at different time post treatment.

<p>Mitochondrial activity and viability in <i>Saprolegnia</i> spores following boric acid (1 g/L) and bronopol (100 mg/L) treatment was compared to non-treated control using the MTS assay. The diagram shows the percent of viable spores relative to the non-treated ones calculated as described in the <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0110343#s2" target="_blank">methodology</a> section. Spores viability was significant reduced (p<0.001) in BA and bronopol treated samples at all time points (4–24 h) relative to non-treated control.</p

Effect of boric acid on the integrity of <i>Saprolegnia</i> spore membranes.

<p>Fluorescence microscopy analysis of Propidium Iodide (PI) uptake by <i>Saprolegnia</i> spores. a) non-treated spores kept in water, were able to germinate and to form hyphae that only flourecent green with SYTO 9 (a1) without PI uptake (a2). b) Boric acid treated spores, neither germinate (b1) nor absorb the PI dye (b2). c) Non-viable, bronopol treated <i>Saprolegnia</i> spores showing uptake of SYTO 9 (c1) and PI dye (c2).</p

Effect of boric acid on <i>Saprolegnia</i> mitochondria using fluorescence microscopy.

<p>Fluorescence microscope image showing the concentration of tetramethyl rhodamine (TMRE) staining in healthy non-treated <i>Saprolegnia</i> hyphae (a1) compared to boric acid treated hyphae (b1) where the depolarized mitochondria exhibit reduced red fluorescence. Figure (a2) is showing ROS level in the non-treated control compared to treated <i>Saprolegnia</i> hyphae (b2). TMRE and ROS staining are merged in a3 and b3.</p

Effect of boric acid on <i>Saprolegnia</i> mitochondria using confocal laser scanning microscopy.

<p>Confocal laser scanning microscopy showing the effect of the boric acid on <i>Saprolegnia</i> spore (a) and hyphal (b) mitochondrial activity using MitoTracker Red. a1) Accumulation of the stain in the non-treated control. Gradual reduction in the number of mitochondria in treated spores 4 (a2), 12 (a3), and 24 (a4) hours after boric acid treatment. b1) <i>Saprolegnia</i> hyphae with densely distributed mitochondria indicating high activity in the non-treated control. Pronounced degradation of hyphal mitochondria 4 (b2), 12 (b3), and 24 (b4) hours post boric acid treatment. <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0110343#pone-0110343-g003" target="_blank">Figure 3 c and d</a> show the average fluorescence intensity of BA treated <i>Saprolegnia</i> spores (c) and hyphae (d) compared to the non-treated control following 24 h exposure.</p

Alterations in <i>Saprolegnia</i> spores following boric acid treatment.

<p>Transmission electron microscopy image of an untreated <i>Saprolegnia</i> spore (a) and a <i>Saprolegnia</i> spore treated with boric acid (1 g/L) for 4 h (b). Normal, well defined mitochondrial structure is seen in the non-treated spores (a1 and a2) compared to the spore that has been exposed to boric acid (b1 and b2). Different degrees of degenerative changes were observed in the mitochondria of the treated spore (circle). The condensed nucleus (N) with disintegrated nuclear membrane is seen in the treated spore (b1), but this was not a consistent finding and seen only in a few spores.</p

Effects of cnidarian biofouling on salmon gill health and development of amoebic gill disease

<div><p>This study examines the potential implications of biofouling management on the development of an infectious disease in Norwegian farmed salmon. The hydroid <i>Ectopleura larynx</i> frequently colonises cage nets at high densities (thousands of colonies per m²) and is released into the water during regular <i>in-situ</i> net cleaning. Contact with the hydroids’ nematocysts has the potential to cause irritation and pathological damage to salmon gills. Amoebic gill disease (AGD), caused by the amoeba <i>Paramoeba perurans</i>, is an increasingly international health challenge in Atlantic salmon farming. AGD often occurs concomitantly with other agents of gill disease. This study used laboratory challenge trials to: (1) characterise the gill pathology resulting from the exposure of salmon to hydroids, and (2) investigate if such exposure can predispose the fish to secondary infections–using <i>P</i>. <i>perurans</i> as an example. Salmon in tanks were exposed either to freshly ‘shredded’ hydroids resembling waste material from net cleaning, or to authentic concentrations of free-living <i>P</i>. <i>perurans</i>, or first to ‘shredded’ hydroids and then to <i>P</i>. <i>perurans</i>. Gill health (AGD gill scores, non-specific gill scores, lamellar thrombi, epithelial hyperplasia) was monitored over 5 weeks and compared to an untreated control group.</p><p>Nematocysts of <i>E</i>. <i>larynx</i> contained in cleaning waste remained active following high-pressure cleaning, resulting in higher non-specific gill scores in salmon up to 1 day after exposure to hydroids. Higher average numbers of gill lamellar thrombi occurred in fish up to 7 days after exposure to hydroids. However, gill lesions caused by hydroids did not affect the infection rates of <i>P</i>. <i>perurans</i> or the disease progression of AGD. This study discusses the negative impacts hydroids and current net cleaning practices can have on gill health and welfare of farmed salmon, highlights existing knowledge gaps and reiterates the need for alternative approaches to net cleaning.</p></div

Scanning electron microscope images of a hydroid tentacle and nematocysts.

<p>a) Close-up of a tentacle of <i>E</i>. <i>larynx</i>, showing cnidocilia of undischarged nematocysts protruding the surface, ready to discharge on contact. b) Two discharged stenotele nematocysts (identified according to Östmann et al. [<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0199842#pone.0199842.ref038" target="_blank">38</a>]) found after triggering release with acetic acid.</p

Hydroids on net sample attached to a frame for high-pressure cleaning.

<p>Hydroids on net sample attached to a frame for high-pressure cleaning.</p

Gill filament of Atlantic salmon showing a lamellar thrombus after exposure to hydroids (H&E and MSB stained).

<p>Gill filament of Atlantic salmon showing a lamellar thrombus after exposure to hydroids (H&E and MSB stained).</p