Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1
DIPTOSH DAS
 -- 2935 2022-04-20 08:04:23 |
2 format correct
Vivi Li
 + 135 word(s) 3070 2022-04-21 03:36:59 |

Video Upload Options

We provide professional Video Production Services to translate complex research into visually appealing presentations. Would you like to try it?
No, upload directly Yes

Confirm

Are you sure to Delete?
Yes No
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Das, D.; Guo, S.; , .; Sabaratnam, V.; Bahkali, A.; Pang, K. Fungi and Climate Change. Encyclopedia. Available online: https://encyclopedia.pub/entry/21975 (accessed on 17 November 2024).
Das D, Guo S,  , Sabaratnam V, Bahkali A, Pang K. Fungi and Climate Change. Encyclopedia. Available at: https://encyclopedia.pub/entry/21975. Accessed November 17, 2024.
Das, Diptosh, Sheng-Yu Guo,  , Vikineswary Sabaratnam, Ali Bahkali, Ka-Lai Pang. "Fungi and Climate Change" Encyclopedia, https://encyclopedia.pub/entry/21975 (accessed November 17, 2024).
Das, D., Guo, S., , ., Sabaratnam, V., Bahkali, A., & Pang, K. (2022, April 20). Fungi and Climate Change. In Encyclopedia. https://encyclopedia.pub/entry/21975
Das, Diptosh, et al. "Fungi and Climate Change." Encyclopedia. Web. 20 April, 2022.
Fungi and Climate Change
Edit
This entry is adapted from the peer-reviewed paper 10.3390/jof8030291

With the over 2000 marine fungi and fungal-like organisms documented so far, some have adapted fully to life in the sea, while some have the ability to tolerate environmental conditions in the marine milieu. These organisms have evolved various mechanisms for growth in the marine environment, especially against salinity gradients. Marine, freshwater and terrestrial fungi and fungal-like organisms vary greatly in their response to salinity. Generally, terrestrial and freshwater fungi grow, germinate and sporulate better at lower salinities, while marine fungi do so over a wide range of salinities. Zoosporic fungal-like organisms are more sensitive to salinity than true fungi, especially Ascomycota and Basidiomycota. Labyrinthulomycota and marine Oomycota are more salinity tolerant than saprolegniaceous organisms in terms of growth and reproduction. Wide adaptability to saline conditions in marine or marine-related habitats requires mechanisms for maintaining accumulation of ions in the vacuoles, the exclusion of high levels of sodium chloride, the maintenance of turgor in the mycelium, optimal growth at alkaline pH, a broad temperature growth range from polar to tropical waters, and growth at depths and often under anoxic conditions, and these properties may allow marine fungi to positively respond to the challenges that climate change will bring.

ocean acidification adaptation deep sea global warming mangrove fungi physiology stress response transcriptome seawater

1. Introduction

The marine ecosystem is host to some 1900 fungi in 769 genera and 133 fungal-like organisms that have evolved for life in the sea (www.marinefungi.org, accessed on 15 December 2021) [1][2]. They include saprobes, parasites and endophytes, and are particularly common in mangroves (500 taxa, [3]) and salt marshes (486 taxa, [4]). This number of marine fungi is low in comparison to the number of terrestrial fungi, but marine fungi are predominantly saprobes and rely on the abundant organic matter available in coastal environments [5]. In the sea, a number of factors affects fungi growth, such as salinity, temperature [6], hydrostatic pressure in the deep-sea [7] and the anoxic conditions of the sediment [8]. Light may affect reproductive behavior of marine fungi [9]. Hydrocarbon [10][11] and plastic [12][13] pollution in the sea may also affect fungal growth behavior. Marine fungi/fungal-like organisms have evolved mechanisms for growth in the marine environment in response to salinity, and these are discussed in this entry. In undertaking this entry, it is important to also consider how terrestrial and freshwater fungi respond to saline conditions in estuaries and mangroves where there are great fluctuations in water salinity.
One of the first studies on the effect of salinity on growth of marine fungi was by Barghoorn and Linder [14], but have researchers made any progress in their understanding of the mechanisms that govern why fungi have been so successful in their colonization and adaptation to marine habitats? Not only do they tolerate high salinity conditions, but they also adapt to life in the sea. Too much time has been spent in trying to define what a marine fungus is, rather than understanding the different requirements of such a diverse fungal community. Perhaps researchers have to accept that not all marine fungi need to necessarily conform to the same physiological requirements in their response to salinity. The current definition of a marine fungus is "any fungus that is recovered repeatedly from marine habitats, because: (1) it is able to grow and/or sporulate (on substrata) in marine environments; (2) it forms symbiotic relationships with other marine organisms; or (3) it is shown to adapt and evolve at the genetic level or be metabolically active in marine environments" [15]. The perennial issue is for those fungi referred to as ‘marine-derived fungi’ often isolated in the search for novel bioactive compounds [15][16] and generally asexual morphs of genera such as Aspergillus, Penicillium and Stanjemonium. Endophytes/endozoans isolated from many plant and animal hosts, and those listed in metabarcoding studies are typical terrestrial taxa [17][18]. It is therefore appropriate to review what determines if a fungus is marine or is simply transient in the marine ecosystem [19].
The response of marine fungi to salinity gradients is of potential importance in terms of climate change with expected high temperatures, increased concentrations of salt, high hydrostatic pressures, and extreme pH. Knowledge of the underlying mechanisms for the adaptation of marine and terrestrial fungi to such events has implications for the biodiversity of marine habitats [20].

2. Growth of Terrestrial and Marine Fungi on Seawater Media

Early studies on marine fungi were dominated by surveys and descriptions of new taxa, their ability to decay wood, and salinity tolerance, especially the requirement for sodium concentrations, as in seawater. Thus, numerous studies have investigated the salinity tolerance of selected marine fungi [14][21][22][23][24][25][26][27][28][29][30][31][32][33][34], to mention but a few. Jones and Jennings [24] warned that a simple comparison of growth of fungi in distilled water and in seawater media does not give a complete picture of the growth of fungi under saline conditions. Here researchers comment on data derived over a range of salinities. A variety of responses have documented the growth of marine fungi, but generally mycelial growth occurred at all salinities with optimum growth varying from 10–50% seawater [24]. Two fungal-like organisms (Althornia crouchi, Ostracoblabe implexa) did not grow in distilled water or in 40% seawater [28]. Most terrestrial and freshwater fungi tested grew better at lower salinities with decreasing growth at higher salinities [22][28], however, a few showed optimum growth under fully saline conditions, especially asexual morphs like Penicillium notatum (100% seawater), and Aspergillus flavus (80% seawater). This tolerance of high salinities may account for why so many such genera are encountered in seawater column samples and in hydrothermal vents [1][35].
Subsequently, experiments were conducted to determine which elements in artificial seawater were tolerated by marine fungi in shake culture. There was no absolute requirement for sodium by Paradendryphiella salina (synonym Dendryphiella salina), but it enhanced growth at low salinities and inhibited growth at higher concentrations [36]. The addition of potassium to a basal medium produced the greatest growth. The bivalent cations magnesium, calcium and strontium inhibited dry weight at all concentrations, but removed sodium inhibition when added at low concentrations (25 M-equiv.). The same pattern was observed in other marine and terrestrial fungi, but varied from species to species. That study indicated that cation content in a medium is critical for the growth of fungi and the permeability of the fungal mycelium [36]. The sodium ion seems to be a key element affecting the growth of marine Chytridiomycota. A species of Phlyctochytrium produced the best growth at 237 mM sodium, and poor growth was observed at 0 mM or concentrations higher than 560 mM [37]. Mg(II) and Ca(II) were essential ions for growth of the same isolate Phlyctochytrium sp. [38].
Some fungi tolerate high concentrations of sodium chloride (NaCl), for example the asexual morph Asteromyces cruciatus tolerated concentrations of 2 M NaCl in liquid media with increased tolerance to 2.5 M NaCl with the addition of 0.05 M CaCl2. Jones and Ward [39] demonstrated that in media with high concentrations of sodium chloride, Asteromyces cruciatus produced septate conidia, rather than the normal unicellular conidia.
Corollospora is a species-rich genus, with most species occurring in association with sand and wood with a worldwide distribution [40][41][42]. Its arenicolous habit means species are exposed to great variations in salinity, especially during the intertidal period. Seventeen arenicolous fungi, ten of which are Corollospora strains, were grown on cornmeal agar in artificial seawater (100%) and at temperatures from 15–40 °C. There was little or no growth at 15 °C and their optimum temperatures are summarised in Table 1. Four Corollospora species were able to grow at 40 °C: C. cinnamomea, C. colossa, C. maritimaand C. pulchella. The effect of salinity and temperature was also investigated with all the fungi exhibiting a positive response to varying salinities (Table 1). Group I are those that exhibited a higher salinity optimum, while Group II fungi had lower salinity optima of 60% and below at most temperatures tested. Four Corollospora maritima strains preferred a high salinity optimum for growth in 80–100% seawater.
Table 1. The optimum salinities (% seawater) for the growth of marine fungi at each temperature investigated. NB: All data used in this table were taken from the linear part of the growth curve. -: no growth; nt: not tested.
Fungi Temperature (°C) Culture in Days
5 °C 15 °C 25 °C 30 °C 35 °C 40 °C
Group I (high salinity optima)              
Arenariomyces trifucatus - 80 80 80 100 - 35
Corollospora besarispora - 80 80 60 80 - 42
Corollospora cinnamomea - 20 100 100 100 100 12
Corollospora colossa - 40 100 80 100 100 30
Corollospora maritima CM1 - - 100 80 80 100 9
Corollospora maritima PP4169 - 80 100 80 - nt 13
Corollospora maritima PP5089 - 20 100 80 100 nt 13
Corollospora maritima PP5197 - 100 100 100 - nt 49
Corollospora novofusca - 20 40 100 100 - 20
Corollospora pulchella - 80 60 60 80 100 15
Corollospora gracilis - 100 80 80 100 nt 13
Savoryella appendiculata - 100 60 80 80 - 48
Torpedospora radiata - 80 60 60 200 - 8
Asteromyces cruciatus 80 80 100 100 - nt 7
Lulworthia crassa - 80 80 60 - - 16
Group II (low salinity optima)              
Carbosphaerella leptosphaerioides - 60 40 60 60 - 28
Corollospora lacera 20 0 20 100 - - 25

3. Fungal-like Organisms and Their Response to Saline Conditions

The fungal-like organisms of the Hyphochytriomycota, Oomycota and Labyrinthulomycota all have marine representatives, especially in mangrove habitats (www.marinefungi.org, accessed on 15 December 2021). Marine representatives of the orders Peronosporales, Pythiales and Saprolegniales (Oomycota) have been widely studied for their growth at various salinities and sodium chloride concentrations (Table 2 and Table 3). They are well adapted to the fluctuating salinities found in mangroves with the production of both asexual and sexual stages [43][44][45].
Table 2. Growth of marine Oomycota and Labyrinthulomycota in various concentrations of sodium chloride.
Species Growth Optimum Remark Reference
Oomycota      
Haliphthoros milfordensis 2.5–3.0% Little or no growth at 0–1.5% [46]
Labyrinthulomycota      
Oblongichytrium multirudimentale 2.5–3.0% No growth at 0% or above 5.0% [47]
Thraustochytrium motivum 2.5–3.0% No growth at 0% or above 5.0% [47]
Thraustochytrium roseum 2.5–5.0% Little or no growth at 0.1–0.5% [47]
Schizochytrium aggregatum 2.5–3.0% Little or no growth at 0.5–1.0% [48]
Table 3. Growth and reproduction of marine Oomycota at various concentration of seawater (‰) (adapted from [44]).
Species Growth Optimum Sporulation Optimum
Halophytophthora avicennae 10–20 (up to 60) 10–30 (none above 35)
Halophytophthora vesicula 15–25 (up to 60) 10–15 (none above 35)
Phytopythium kandeliae 10–35 (none above 35) 15–35 (none above 35)
Salispina lobata 20–40 (none above 40) 30–40 (none above 40)
Salisapilia masteri 20–35 (up to 60) 30 (none above 40)
Few members of the Saprolegniaceae have been reported from marine habitats [49], and Harrison and Jones [50] questioned if this was due to their inability to tolerate high salinity levels. Using zoospore suspensions of 17 saprolegniaceous species, they investigated their ability to reproduce asexually at salinities of 0–40% seawater. Normal sporangial development occurred in freshwater, while in 10% seawater only nine produced normal zoospores: e.g., Achlya bisexualis, Protoachyla paradoxa, Thraustyotheca clavata, Saprolegnia parasiticaand a Saprolgenia sp. In three Isoachlya spp. and Achyla raceomsa, some cytoplasmic cleavage occurred. In 20% seawater only small sporangial primordia were formed, with only S. parasitica and T. clavata producing zoospores. Formation of sexual reproduction was also investigated for the same taxa with most species forming oospheres, but in I. toruloides there was no cleavage of the oogonial cytoplasm. At 20% seawater only P. paradoxa produced mature oospheres, while at salinities above 20%, sexual reproduction was suppressed. Similar results were reported by Höhnk [51][52] for a Saprolegnia sp. when 7.09% salinity inhibited asexual reproduction, although excellent vegetative growth occurred. Clearly members of the Saprolegniales are not well adapted for survival at the higher salinities found in the ocean.
Species of the oomycetous genus Halophytophthora have wide salinity growth ranges. Isolates of H. avicenniae and H. batemanensis were able to grow at 4‰, 8‰, 16‰ and 32‰, although pH and incubation temperature had a combinatorial effect on growth [45]. The wide salinity growth ranges of Halophytophthora isolates suggest that they are well adapted to the salinity variation daily (high/low tide) and seasonally (summer/winter, rainy/dry seasons) in mangrove habitats.

4. Physiological Response to Salinity

This is a topic that has been widely researched by David Jennings and his students [53][54][55][56][57][58][59][60][61] who have carried out extensive studies on marine fungi and their ability to tolerate saline conditions in the marine environment. Do marine fungi require sodium chloride for growth and what are the mechanisms that control osmotic pressure within their mycelium? In an effort to understand these phenomena, a wide range of techniques have been applied to elucidate the mechanisms involved. Jennings [53] highlighted three physiological issues facing a fungus in the marine environment: seawater (1) has a relatively low water potential, (2) contains a relatively high concentration of ions, and (3) has an alkaline pH.
Studies with Paradendryphiella salina showed that sodium stimulates its growth at low salinities but is toxic at high concentrations [36]. This inhibition could be overcome by the addition of magnesium, calcium, strontium and barium (in order of effectiveness), and this was also demonstrated for other marine fungi [36]. The study also demonstrated that the key issue for the growth of the fungus was the permeability of the mycelium to potassium in a high salt medium. This study led to the exploration of a number of factors pertinent to understanding the mechanisms of salt tolerance in marine and other fungi [53]. These included plasma membrane permeability, accumulation of ions in the vacuoles, and the role of polyols in maintaining turgor in the mycelium.
In seawater, how does Paradendryphiella salina control the movement of nutrients and ions into the mycelium? It has already been shown that calcium and other bivalent cations play a role in the movement of potassium and exclusion of sodium from the mycelium. Jones and Jennings [36] proposed that there was a sodium pump for its exclusion from the mycelium. Galpin and Jennings [62] reported the involvement of ATPase in maintaining the K+/Na+ ratio within the mycelium of P. salina, and ATPase is required for the active transport of cations. When P. salina was grown at high salinity and pH, there was an increased activity of membrane-bound ATPase and this aids in good potassium/sodium balance in the cytoplasm [53]. Thus, ATPase is required for active transport of cations and fulfilled by glycolysis.
These studies continued by exploring the accumulation of ions in fungal vacuoles [56][60][63], plasma-membrane permeability [58][64] and the role of polyols in maintaining turgor in the marine fungal mycelium [61][63]. Ions contributed some 60% of the solute potential in the 48-h old mycelium of P. salina grown in the presence of high concentrations of sodium chloride, while polyols contributed 30% [63]. Jennings and Austin [60] showed the importance of mannitol and arabitol in maintaining the total in vivo carbohydrate content of mycelium of P. salina which is required for making optimum turgor for growth. Mannitol and arabitol synthesis within the hypha increased with increasing salinity because these sugar alcohols play the main role in maintaining osmotic pressure as well as correct differential water potential in the mycelium [54]. This was confirmed by Wethered et al. [63], who provided evidence that the polyol content of the mycelium increased with salinity. Holligan and Jennings [58] proposed two pathways of mannitol synthesis; one is directly from glucose entering the hyphae and another is from hexose phosphate derived from pentose phosphate pathway with ATPase hydrolysis (Figure 1A,B). Arabitol synthesis depends upon a stimulation of the pentose phosphate pathway and is derived from pentose sugar (xylulose and ribulose) via the pentose phosphate pathway (Figure 1C). Jennings [56] concluded that mannitol, arabitol, glycerol, and erythritol are the major polyols which are accumulated by mycelium, and variation in carbon sources, such as glucose and fructose, has an effect in the accumulation of various polyols.
Figure 1. (A). Mannitol synthesis from glucose. (B). Mannitol synthesis from the hexose phosphate derived from the pentose phosphate pathway. (C). Arabitol synthesis from pentose sugar via the pentose phosphate pathway.
In contrast to the mycelial fungi discussed above, there is evidence that zoosporic genera in the Thraustochytriales require sodium chloride for growth. Siegenthaler et al. [65][66] suggested that phosphate uptake in Thraustochytrium roseum required sodium chloride. Also, they demonstrated that the presence of the amino acid proline in their cells, as well as high levels of inorganic ions which contribute to the solute potential of the cells. Wethered and Jennings [67] noted that proline concentrations in cells increased with the increased salinity of the medium.
Norkrans [68] and Norkrans and Kylin [69] drew attention to marine occurring yeasts that are halotolerant with growth in the range of 0–24% sodium chloride. Gustafsson and Norkrans [70] and Adler and Gustafsson [71] reported that polyols accumulated in the marine occurring yeast Debaryomyces hansenii due to salt-stress, while Adler [72] showed that the accumulation of glycerol in D. hansenii played a role in osmoregulation.
Fungi in man-made salterns, soda lakes, coastal lagoons and the Dead Sea, tolerate very high environmental NaCl concentrations when compared to the marine fungi discussed here [73][74]. Larsen [73] characterised these fungi into four categories depending on tolerance to NaCl concentrations: non-tolerant up to 1%, slight 10%, moderate 20% and extreme 30% NaCl. These fungi too face similar physiological conditions to marine fungi, namely an external environment with relatively low water potential and high concentration of ions [75]. It is speculated that cation transporters prevent intracellular accumulation of Na+, which would be toxic but plays a role in maintaining the high K+/Na+ ratio required for growth in an environment with high salt content. The halophilic Wallemia ichthyophaga accumulates glycerol, while Hortaea werneckii also accumulates erythritol, arabitol and mannitol as solutes [76][77]. The same mechanism applies to the growth of marine fungi in seawater [53].

5. Marine Fungi and Climate Change

Kumar et al. [78] opined on the ecology and evolution of marine fungi and their potential adaptation to climate change, but did not consider their physiology and tolerance of saline conditions. Marine fungi are unique with many characteristics that define their life in a saline environment. These include wide adaptability to saline conditions in mangroves/estuaries and salterns, mechanisms for maintaining accumulation of ions in the vacuoles, exclusion of high level of sodium chloride, maintaining turgor in the mycelium, optimal growth at alkaline pH, a broad temperature growth range from polar waters to higher temperatures in sand dunes/intertidal periods (0–40 °C), growth at depths and often under anoxic conditions [79]. With these features, marine fungi may well positively respond to the challenges that climate change will bring. Key amongst these will be an increase in CO2 levels, the predicted rise in temperatures, changes (dilution due to melting of the ice caps) in the salinity of seawater and rising sea-levels which will affect the distribution of sea grasses and mangrove/salt marsh plants [78].
1. An increase in CO2 levels will affect the acidity of seawater, which has implications for the growth of fungi with an alkaline pH requirement. Caldeira and Wickett [80] noted that seawater pH has dropped by 0.1 units, and may decrease by a further 0.7 units within the next three centuries. Krause et al. [81] carried out acidification experiments in microcosms with seawater from the Baltic Sea and recorded fungal abundance (as colony forming units). Their results suggested that even moderate acidification may lead to an increase in fungal abundance of almost an order of magnitude. 
2. Marine fungi have a broad tolerance to variation in salinity in terms of mycelial growth, spore germination and sporulation, and therefore should adapt to changes in oceanic salinity (see Figure 2).
Figure 1. Effect of salinity on the production of perithecia, asci and ascospores of fungi.
3. Marine fungi appear to tolerate a wide range in seawater temperature (See Table 1). Although marine fungi are worldwide in distribution, certain taxa may be restricted geographically to the tropics, subtropics, temperate or polar waters [82][83][84][85][86]. However, there is little overlap in fungal species from tropical and temperate regions [87]. Consequently, many marine fungi in temperate regions will have to adapt to increased temperature. Pang et al. [88] have shown that an Aspergillus terreus strain isolated from a shallow hydrothermal vent was able to grow at 45 °C, pH 3, and 30% salinity.

References

  1. Jones, E.B.G.; Pang, K.L.; Abdel-Wahab, M.A.; Scholz, B.; Hyde, K.D.; Boekhout, T.; Ebel, R.; Rateb, M.E.; Henderson, L.; Sakayaroj, J. An online resource for marine fungi. Fungal Divers. 2019, 96, 347–433.
  2. Pang, K.L.; Hassett, B.T.; Shaumi, A.; Guo, S.Y.; Sakayaroj, J.; Chiang, M.W.L.; Yang, C.H.; Jones, E.B.G. Pathogenic fungi of marine animals: A taxonomic perspective. Fungal Biol. Rev. 2021, 38, 92–106.
  3. Devadatha, B.; Jones, E.B.G.; Pang, K.L.; Abdel-Wahab, M.A.; Hyde, K.D.; Sakayaroj, J.; Bahkali, A.H.; Calabon, M.S.; Sarma, V.V.; Suetrong, S.; et al. Occurrence and geographical distribution of mangrove fungi. Fungal Divers. 2021, 106, 137–227.
  4. Calabon, M.S.; Jones, E.B.G.; Promputtha, I.; Hyde, K.D. Fungal biodiversity in salt marsh ecosystems. J. Fungi 2021, 7, 648.
  5. Jones, E.B.G.; Suetrong, S.; Sakayaroj, J.; Bahkali, A.H.; Abdel-Wahab, M.A.; Boekhout, T.; Pang, K.-L. Classification of marine Ascomycota, Basidiomycota, Blastocladiomycota and Chytridiomycota. Fungal Divers. 2015, 73, 1–72.
  6. Jones, E.B.G. Marine fungi: Some factors influencing biodiversity. Fungal Divers. 2000, 4, 53–73.
  7. Burgaud, G.; Hué, N.T.M.; Arzur, D.; Coton, M.; Perrier-Cornet, J.-M.; Jebbar, M.; Barbier, G. Effects of hydrostatic pressure on yeasts isolated from deep-sea hydrothermal vents. Res. Microbiol. 2015, 166, 700–709.
  8. Zain Ul Arifeen, M.; Chu, C.; Yang, X.; Liu, J.; Huang, X.; Ma, Y.; Liu, X.; Xue, Y.; Liu, C. The anaerobic survival mechanism of Schizophyllum commune 20R-7-F01, isolated from deep sediment 2 km below the seafloor. Environ. Microbiol. 2021, 23, 1174–1185.
  9. Cai, M.; Fang, Z.; Niu, C.; Zhou, X.; Zhang, Y. Light regulation on growth, development, and secondary metabolism of marine-derived filamentous fungi. Folia Microbiol. 2013, 58, 537–546.
  10. Bovio, E.; Gnavi, G.; Prigione, V.; Spina, F.; Denaro, R.; Yakimov, M.; Calogero, R.; Crisafi, F.; Varese, G.C. The culturable mycobiota of a Mediterranean marine site after an oil spill: Isolation, identification and potential application in bioremediation. Sci. Total Environ. 2017, 576, 310–318.
  11. Dell’Anno, F.; Rastelli, E.; Sansone, C.; Brunet, C.; Ianora, A.; Dell’Anno, A. Bacteria, fungi and microalgae for the bioremediation of marine sediments contaminated by petroleum hydrocarbons in the omics era. Microorganisms 2021, 9, 1695.
  12. De Tender, C.; Devriese, L.I.; Haegeman, A.; Maes, S.; Vangeyte, J.r.; Cattrijsse, A.; Dawyndt, P.; Ruttink, T. Temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea. Environ. Sci. Technol. 2017, 51, 7350–7360.
  13. Lacerda, A.L.d.F.; Proietti, M.C.; Secchi, E.R.; Taylor, J.D. Diverse groups of fungi are associated with plastics in the surface waters of the Western South Atlantic and the Antarctic Peninsula. Mol. Ecol. 2020, 29, 1903–1918.
  14. Barghoorn, E.S.; Linder, D.H. Marine fungi: Their taxonomy and biology. Farlowia 1944, 1, 395–467.
  15. Pang, K.L.; Overy, D.P.; Jones, E.B.G.; Calado, M.D.; Burgaud, G.; Walker, A.K.; Johnson, J.A.; Kerr, R.G.; Cha, H.J.; Bills, G.F. ‘Marine fungi’ and ‘marine-derived fungi’ in natural product chemistry research: Toward a new consensual definition. Fungal Biol. Rev. 2016, 30, 163–175.
  16. Trisuwan, K.; Rukachaisirikul, V.; Sukpondma, Y.; Phongpaichit, S.; Preedanon, S.; Sakayaroj, J. Lactone derivatives from the marine-derived fungus Penicillium sp. PSU-F44. Chem. Pharm. Bull. 2009, 57, 1100–1102.
  17. Chaeprasert, S.; Piapukiew, J.; Whalley, A.J.; Sihanonth, P. Endophytic fungi from mangrove plant species of Thailand: Their antimicrobial and anticancer potentials. Bot. Mar. 2010, 53, 555–564.
  18. Vargas-Gastélum, L.; Riquelme, M. The mycobiota of the deep sea: What omics can offer. Life 2020, 10, 292.
  19. Park, D. On the ecology of heterotrophic microorganisms in freshwater. Trans. Br. Mycol. Soc. 1972, 58, 291–299.
  20. Velez, P. Impact of salinity stress on growth and development of aquatic fungi. In Microorganisms in Saline Environments: Strategies and Functions; Giri, B., Varma, A., Eds.; Springer: Cham, Germany, 2019; pp. 155–168.
  21. Ritchie, D. Salinity optima for marine fungi affected by temperature. Am. J. Bot. 1957, 44, 870–874.
  22. Gray, W.D.; Pinto, P.V.C.; Pathak, S.G. Growth of fungi in sea water medium. Appl. Microbiol. 1963, 11, 501–505.
  23. Kirk, P.W., Jr. Morphogenesis and microscopic cytochemistry of marine pyrenomycete ascospores. Nova Hedw. Beih. 1966, 22, 1–128.
  24. Jones, E.B.G.; Jennings, D.H. The effect of salinity on the growth of marine fungi in comparison with non-marine species. Trans. Br. Mycol. Soc. 1964, 47, 619–625.
  25. Meyers, S.P.; Simms, J. Thalassiomycetes VI. Comparative growth studies of Lindra thalassiae and lignicolous Ascomycete species. Can. J. Bot. 1965, 43, 379–392.
  26. Meyers, S.P.; Hoyo, L. Observations on the growth of the marine Hyphomycete Varicosporina ramulosa. Can. J. Bot. 1966, 44, 1133–1140.
  27. Meyers, S.P.; Scott, E.; Thalassiomycetes, X. Variation in growth and reproduction of two isolates of Corollospora maritima. Mycologia 1967, 59, 446–455.
  28. Jones, E.B.G.; Byrne, P.J.; Alderman, D.J. The response of fungi to salinity. Vie Milieu Suppl. 1971, 22, 265–280.
  29. Davidson, D.E. The effect of salinity on a marine and a freshwater Ascomycete. Can. J. Bot. 1974, 52, 553–563.
  30. Byrne, P.J.; Jones, E.B.G. Effect of salinity on spore germination of terrestrial and marine fungi. Trans. Brit. Mycol. Soc. 1975, 64, 497–503.
  31. Byrne, P.J.; Jones, E.B.G. Effect of salinity on the reproduction of terrestrial and marine fungi. Trans. Brit. Mycol. Soc. 1975, 65, 185–200.
  32. Gessner, R.V. In vitro growth and nutrition of Buergenerula spartinae, a fungus associated with Spartina alterniflora. Mycologia 1976, 68, 583–599.
  33. Curran, P. The effect of temperature, pH, light and dark on the growth of fungi from Irish coastal waters. Mycologia 1980, 72, 350–358.
  34. Crabtree, S.L.; Gessner, R.V. Growth and nutrition of the salt marsh fungi Pleospora gaudifroyi and Camarosporium roumeguerii. Mycologia 1982, 74, 640–647.
  35. Richards, T.A.; Leonard, G.; Mahé, F.; del Campo, J.; Romac, S.; Jones, M.D.M.; Maguire, F.; Dunthorn, M.; De Vargas, C.; Massana, R.; et al. Molecular diversity and distribution of marine fungi across 130 European environmental samples. Proc. R. Soc. B 2015, 282, 20152243.
  36. Jones, E.B.G.; Jennings, D.H. The effect of cations on the growth of fungi. New Phytol. 1965, 64, 86–100.
  37. Amon, J.P. An estuarine species of Phlyctochytrium (Chytridiales) having a transient requirement for sodium. Mycologia 1976, 68, 470–480.
  38. Amon, J.P.; Arthur, R.D. Nutritional studies of a marine Phlyctochytrium sp. Mycologia 1981, 73, 1049–1055.
  39. Jones, E.B.G.; Ward, A.W. Septate conidia in Asteromyces cruciatus. Trans. Brit. Mycol. Soc. 1973, 61, 181–186.
  40. Nakagiri, A.; Tubaki, K. Ascocarp peridial wall structure in Corollospora and allied genera of Halosphaeriaceae. In The Biology of Marine Fungi; Moss, S.T., Ed.; Cambridge University Press: Cambridge, UK, 1986; pp. 245–251.
  41. Lintott, W.H.; Lintott, E.A. Marine fungi from New Zealand. In Fungi in Marine Environments; Hyde, K.D., Ed.; Fungal Diversity Research Series: Hong Kong, China, 2002; pp. 285–292.
  42. Velez, P.; Alejandri-Ramírez, N.D.; González, M.C.; Estrada, K.J.; Sanchez-Flores, A.; Dinkova, T.D. Comparative transcriptome analysis of the cosmopolitan marine fungus Corollospora maritima under two physiological conditions. G3 Genes Genomes Genet. 2015, 5, 1805–1814.
  43. Leaño, E.M.; Vrijmoed, L.L.P.; Jones, E.B.G. Physiological studies on Halophytophthora vesicula (straminipilous fungi) isolated from fallen mangrove leaves from Mai Po, Hong Kong. Bot. Mar. 1998, 41, 411–419.
  44. Leaño, E.M.; Jones, E.B.G.; Vrijmoed, L.L.P. Why are Halophytophthora species well adapted to mangrove habitats? Fungal Divers. 2000, 5, 131–151.
  45. Su, C.J.; Hsieh, S.Y.; Chiang, M.W.L.; Pang, K.L. Salinity, pH and temperature growth ranges of Halophytophthora isolates suggest their physiological adaptations to mangrove environments. Mycology 2020, 11, 256–262.
  46. Vishniac, H.S. A new marine Phycomycete. Mycologia 1958, 50, 66–79.
  47. Goldstein, S. Development and nutrition of new species of Thraustochytrium. Am. J. Bot. 1963, 50, 271–279.
  48. Goldstein, S.; Belsky, M. Axenic culture studies of a new marine phycomycete possessing an unusual type of asexual reproduction. Am. J. Bot. 1964, 51, 72–78.
  49. Johnson, T.W.; Sparrow, F.K. Fungi in Oceans and Estuaries; Cramer: Weinheim, Germany, 1961; p. 668.
  50. Harrison, J.L.; Jones, E.B.G. Effect of salinity on sexual and asexual sporulation of members of the Saprolegniaceae. Trans. Br. Mycol. Soc. 1975, 65, 389–394.
  51. Höhnk, W. Ein Beitrag zur Kenntnis der Phycomyceten des Brackwassers. Kiel. Meeresforsch. 1939, 3, 337–361.
  52. Höhnk, W. Eine neue uferbewohnende Saprolegniazee: Calyptralegnia ripariensis nov. spec. Veröffentlichungen Des Inst. Für Meeresforsch. Bremerhav. 1953, 2, 230–235.
  53. Jennings, D.H. Fungal growth in the sea. In The Biology of Marine Fungi; Moss, S.T., Ed.; Cambridge University Press: Cambridge, UK, 1986; pp. 1–10.
  54. Jennings, D.H. Cations and filamentous fungi: Invasion of the sea and hyphal functioning. In Ion Transport in Plants; Anderson, W.P., Ed.; Academic Press: London, UK, 1973; pp. 323–335.
  55. Jennings, D.H. Some aspects of the physiology and biochemistry of marine fungi. Biol. Rev. 1983, 58, 423–459.
  56. Jennings, D.H. Polyol metabolism in fungi. Adv. Microb. Physiol. 1985, 25, 149–193.
  57. Holligan, P.M.; Jennings, D.H. Carbohydrate metabolism in the fungus Dendryphiella salina. I. Changes in the Levels of Soluble Carbohydrates during Growth. New Phytol. 1972, 71, 569–582.
  58. Holligan, P.M.; Jennings, D.H. Carbohydrate metabolism in the fungus Dendryphiella salina. II. The influence of different carbon and nitrogen sources on the accumulation of mannitol and arabitol. New Phytol. 1972, 71, 583–594.
  59. Holligan, P.M.; Jennings, D.H. Carbohydrate metabolism in the fungus Dendryphiella salina. III. The effect of the nitrogen source on the metabolism of -and -glucose. New Phytol. 1972, 71, 1119–1133.
  60. Jennings, D.H.; Austin, S. The stimulatory effect of the non-metabolisable sugar 3-O-methyl glucose on the conversion of mannitol and arabitol to polysaccharide and other insoluble compounds in the fungus Dendryphiella salina. J. Gen. Microbiol. 1973, 75, 287–294.
  61. Lowe, D.A.; Jennings, D.H. Carbohydrate metabolism in the fungus Dendryphiella salina. V. The pattern of label in arabitol and polysaccharide after growth in the presence of specifically labelled carbon sources. New Phytol. 1975, 74, 67–79.
  62. Galpin, M.F.J.; Jennings, D.H. Histochemical study of the hyphae and the distribution of adenosine triphosphatase in Dendryphiella salina. Trans. Brit. Mycol. Soc. 1975, 65, 477–483.
  63. Wethered, J.M.; Metcalf, E.C.; Jennings, D.H. Carbohydrate metabolism in the fungus Dendryphiella salina. VIII. The contribution of polyols and ions to the mycelial solute potential in relation to the external osmoticum. New Phytol. 1985, 101, 631–649.
  64. Galpin, M.F.J.; Jennings, D.H. A plasma-membrane ATPase from Dendryphiella salina: Cation specificity and interaction with fusicoccin and cyclic AMP. Trans. Br. Mycol. Soc. 1980, 75, 35–46.
  65. Siegenthaler, P.A.; Belsky, M.M.; Goldstein, S. Phosphate uptake in an obligately marine fungus: A specific requirement for sodium. Science 1967, 155, 93–94.
  66. Siegenthaler, P.A.; Belsky, M.M.; Goldstein, S.; Menna, M. Phosphate uptake in an obligately marine fungus. II. Role of culture conditions, energy sources and inhibitors. J. Bacteriol. 1967, 93, 1281–1288.
  67. Wethered, J.M.; Jennings, D.H. The major solutes contributing to the solute potential of Thraustochytrium aureum and T. roseum after growth in media of different salinities. Trans. Br. Myol. Soc. 1985, 85, 439–446.
  68. Norkrans, B. Studies on marine occurring yeasts. Growth related to pH, NaCl concentrations and temperature. Arch. Mikrobiol. 1966, 54, 374–392.
  69. Norkrans, B.; Kylin, A. Regulation of potassium to sodium and of the osmotic potential in relation to salt tolerance in yeasts. J. Bacteriol. 1969, 100, 836–845.
  70. Gustafsson, L.; Norkrans, B. On the mechanism of salt tolerance. Production of glycerol and heat during growth of Debaryomyces hansenii. Arch. Microbiol. 1976, 110, 177–183.
  71. Adler, L.; Gustafsson, L. Polyhydric alcohol production and intracellular amino acid pool in relation to halotolerance of the yeast Debaryomyces hansenii. Arch. Microbiol. 1980, 124, 123–130.
  72. Adler, L. Physiological and biochemical characteristics of the yeast Debaryomyces hansenii in relation to salinity. In The Biology of Marine Fungi; Moss, S.T., Ed.; Cambridge University Press: Cambridge, UK, 1986; pp. 81–89.
  73. Larsen, H. Halophilic and halotolerant microorganisms- an overview and historical perspective. FEMS Microbiol. Rev. 1986, 39, 3–7.
  74. Kis-Papo, T.; Oren, A.; Wasser, S.P.; Nevo, E. Survival of filamentous fungi in hypersaline Dead Sea water. Microb. Ecol. 2003, 45, 183–190.
  75. Gunde-Cimerman, N.; Ramos, J.; Plemenitas, A. Halotolerant and halophilic fungi. Mycol. Res. 2009, 113, 1231–1241.
  76. Gunde-Cimerman, N.; Butinar, L.; Sonjak, S.; Turk, M.; Uršič, V.; Zalar, P.; Plemenitaš, A. halotolerant and halophilic fungi from coastal environments in the Arctics. In Adaptation to Life at High Salt Concentrations in Archaea, Bacteria, and Eukarya (Cellular Origin, Life in Extreme Habitats and Astrobiology); Gunde-Cimerman, N., Oren, A., Plemenitaš, A., Eds.; Springer: Dordrecht, The Netherlands, 2005; pp. 397–423.
  77. Zajc, J.; Zalar, P.; Plemenitaš, A.; Gunde-Cimerman, N. The mycobiota of the salterns. In Biology of Marine Fungi; Raghukumar, C., Ed.; Springer-Verlag: Berlin/Heidelberg, Germany, 2012; pp. 133–158.
  78. Kumar, V.; Sarma, V.V.; Thambugala, K.M.; Huang, J.J.; Li, X.Y.; Hao, G.F. Ecology and evolution of marine fungi with their adaptation to climate change. Front. Microbiol. 2021, 12, 719000.
  79. Raghukumar, S. Origin and Evolution of Marine Fungi. In Fungi in Coastal and Oceanic Marine Ecosystems; Raghukumar, S., Ed.; Springer International Publishing AG: Cham, Germany, 2017; pp. 307–321.
  80. Caldeira, K.; Wickett, M.E. Anthropogenic carbon and ocean pH. Nature 2003, 425, 365.
  81. Krause, E.; Wichels, A.; Gimenez, L.; Gerdts, G. Marine fungi may benefit from ocean acidification. Aquat. Microb. Ecol. 2013, 69, 59–67.
  82. Hughes, G.C. Biogeography and the marine fungi. In The Biology of Marine Fungi; Moss, S.T., Ed.; Cambridge University Press: Cambridge, UK, 1986; pp. 275–295.
  83. Hughes, G.C. Geographical distribution of the higher marine fungi. Veroff. Inst. Meeresforsch. Bremerh. Suppl. 1974, 5, 419–441.
  84. Hyde, K.D. Frequency of occurrence of lignicolous marine fungi in the tropics. In The Biology of Marine Fungi; Moss, S.T., Ed.; Cambridge University Press: Cambridge, UK, 1986; pp. 311–322.
  85. Hyde, K.D.; Jones, E.B.G. Marine mangrove fungi. Mar. Ecol. 1988, 9, 15–33.
  86. Schmit, J.P.; Shearer, C.A. A checklist of mangrove associated fungi, their geographical distribution and known host plants. Mycotaxon 2003, 85, 423–477.
  87. Jones, E.B.G.; Pang, K.L. Tropical aquatic fungi. Biodivers. Conserv. 2012, 21, 2403–2423.
  88. Pang, K.L.; Chiang, M.W.; Guo, S.Y.; Shih, C.Y.; Dahms, H.U.; Hwang, J.S.; Cha, H.J. Growth study under combined effects of temperature, pH and salinity and transcriptome analysis revealed adaptations of Aspergillus terreus NTOU4989 to the extreme conditions at Kueishan Island Hydrothermal Vent Field, Taiwan. PLoS ONE 2020, 15, e0233621.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
Upload a video for this entry
Information
Subjects: Biology
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
DIPTOSH DAS
,
Sheng-Yu Guo
,
,
Vikineswary Sabaratnam
,
Ali Bahkali
,
Ka-Lai Pang
View Times: 636
Revisions: 2 times (View History)
Update Date: 21 Apr 2022
Table of Contents
    1000/1000
    Hot Most Recent
    ScholarVision Creations
    Feedback