Number of recorded fauna of Bangladesh.
\r\n\tNatural hazards are potentially damaging physical events and phenomena, which may cause the loss of life, injury or human life disruption, property damage, social, economic, and political disruption, or environmental degradation.
\r\n\tNatural hazards can be divided into different groups: geological, hydro-meteorological, climatological, outer space, and biological hazards.
\r\n\tA disaster is a serious disruption of the normal functioning of a society causing widespread human, material, economic or environmental losses. A disaster results from the combination of hazards, conditions of vulnerability and insufficient capacity or measures to reduce the potential negative consequences of risk, and exposure.
\r\n\tEarthquakes, volcano eruptions, tsunamis, curst, suffusion, coast erosion, and landslides belong to geological hazards.
\r\n\tHydro-meteorological and climatological hazards are the most frequent causes of the disaster events among all natural hazards. The most common meteorological hazards are heavy rains, tornadoes, storms, hurricanes, droughts, tropical cyclones, rainstorm floods, heat waves, and low-temperature disasters.
\r\n\tA comparison of the loss events and fatalities shows that the regions with economically less-developed countries have more fatalities, but more rich countries have higher damage during disasters.
\r\n\tThe Book addresses principles, concepts and paradigms of environmental economics connected discipline, as well as operational terms, materials, tools, techniques, and methods including processes, procedures and implications.
\r\n\tThe Book equips professionals and others with a formal understanding of environmental economics topics. Clarifies the similarities or differences in fundamental concepts and principles in the discipline. Captures the wide range of expanding disciplinary activities under a single umbrella of environmental economics concept.
Grasses, shrubs, and different palatable parts of trees, i.e., leaves, flowers, fruits, and seeds, that have nutritive values constitute fodder of wild herbivores [1]. The wildlife population greatly depends on the habitat richness with food, nesting, and breeding environment. The wild fruit and fodder-producing plants play a great role in maintaining ecosystem food supply. The plants are from different habit forms and taxonomic families. Plants from Poaceae, Cyperaceae, Fabaceae, Moraceae, Myrtaceae, and Zingiberaceae families particularly the leguminous plants dominate the fodder-producing plants [2]. There are significant seasonal variation of fodder availability and composition to which the wild animals’ nutrition needs are naturally adapted [3]. Insects, birds, chordates, and reptiles have different nesting and breeding natures which varies widely from each other. The habitat resources and overall conditions including food, water, shade, nesting, etc. are needed by a species for survival and reproductive success [4]. Moreover, habitat is organism-specific; the appropriate combination of necessary abiotic and biotic components for successful reproduction and survival varies by species [5].
\nThe global forests are drastically shrinking day by day due to a huge pressure on forests for conversion to other land use, human settlement, excessive resource extraction, etc. All these are affecting biodiversity negatively particularly the wildlife habitat which is degrading in an alarming way which leads to rapid shrinking of wildlife population and making them threatened. Declaring protected area (PA) is a worldwide strategy accepted for wildlife habitat conservation and ensuring undisturbed breeding ground by halting further fragmentation and degradation of habitat. This chapter presents the status of the protected areas from different corners of Bangladesh in terms of fodder yielding plant composition as well as the importance of fodder plant management for maintaining a healthy wildlife habitat. We identified three protected areas of characteristic features for studying the wild fodder yielding plants so that these represent all the PA of Bangladesh. This chapter also presents a brief account on the protected area management in Bangladesh as well as potentiality of those areas as wildlife habitats.
\nQuality of the wildlife habitat largely depends on the nature and composition of vegetation as it determines the nesting, breeding, and feeding potentiality of the habitats. The habitat degradation is causing loss of regeneration of many of the fodder plants. Moreover, overexploitation is also responsible for sharp reduction of the population size of the fodder plants [6]. It is important to detect the food habit of the herbivores and identify the fodder plants and their interactions with animals for sustainable management of the protected areas through wildlife conservation and undisturbed breeding ground [7]. But, in Bangladesh, there is a great dearth of information and research on the wild fodder-bearing trees. Information on the status and availability of plants will be helpful for better planning and management interventions of the PAs as wildlife habitats. The plants used as fodder by the animals are rich in necessary nutrition, i.e., protein, carbohydrate, fiber, etc. The ability of fodder plants to provide this range of nutrients is considered to evaluate potentiality of their nutritive values. Considering the mentioned situations, the study was undertaken to assess the overall composition of fodder plants as well as highlight their contributions for better maintaining a wildlife habitat. However, there is still a need for investigation of the nutritive values as many of the species were not explored yet [6]. We believe this study will fill up the knowledge gap on fodder yielding plant composition in the PA of Bangladesh as well as it will show the way for further research and interventions to habitat conservation and sustainable management.
\nBangladesh is the home of more than 3883 species of lower faunal groups along with 653 fish species, 49 amphibians, 154 reptiles, 706 birds, and 128 mammals. The fish communities including both freshwater and marine species are so diverse that they account an astonishing 3% of the world’s total fish species. In addition to the 383 resident birds, there are 323 migratory birds which visit our country especially during the winter. Both of these represent an amazing 7.2% of the world’s total bird species. Mammals constitute 2.28% of the world’s mammal species among which seven are marine in nature [8]. Though our wildlife diversity is very rich, but over the last century unfortunately 13 species have become extinct from Bangladesh. Due to continuity of habitat degradation, many more are on the brink of extinction. Different reports indicated that 23% of vertebrates of Bangladesh are facing different levels of threats which are increasing exponentially with rapid habitat destruction. The situation is even more grim for the 57% of reptiles and 36% of mammals which are facing different levels of threats in our country [9]. Recently, the IUCN listed 40 species of mammals, 41 aves, 58 reptiles, and 8 amphibians that are struggling under various degrees of threat of extinction. It is obvious that the present wildlife population is confined and distributed irregularly in limited forest patches of Bangladesh (Table 1).
\n\nAmong the 5 global ecological domains and 20 global ecological zones of the world, 33% of Bangladesh belongs to tropical rain forest GEZ and 67% to tropical moist deciduous forest GEZ of the tropical domains of global ecological domains [10]. The variation in climatic features, i.e., temperature, rainfall, soil, and hydrology, led to the development of 25 bioecological zones with distinct characteristics. Bangladesh has 1.45 million ha of forest land (9.8% of total area) of which 1.21 million ha (84% of forest) is natural forest and 0.24 million ha (16% of forest) is plantations [10].
\nVegetation characteristics divided the natural forests of Bangladesh into evergreen/semievergreen, deciduous, and mangrove forest. Noncontinuous freshwater swamp is distributed in the northeast basin. Tropical evergreen and semievergreen forests constituting 44% of natural forest are extended over Chittagong, Chittagong Hill Tracts (CHT), Cox’s Bazar, and Sylhet covering an area of 6700 km2 which is about 4.54% of total landmass of Bangladesh. Dominant native plant species include species of Dipterocarpus spp., Artocarpus spp., Ficus spp., Syzygium spp., Mangifera spp., Tectona grandis, etc. The moist deciduous Sal forest of Bangladesh is mainly consisted of Madhupur tract which is located in the central part covering an area of 340 km2 [10]. Dominant trees of this forest are Shorea robusta, Lagerstroemia speciosa, Dillenia pentagyna, Adina cordifolia, Terminalia spp., Albizia spp., etc. The Sundarbans, the largest single patch of mangrove forest, is located at the southern extremity of the Padma (Ganges) and Jamuna (Brahmaputra) delta which covers about 5770 km2 area [11]. Fairly dense evergreen plant species of 10–15 m height is the main feature of this forest. These species are adapted for living under saline condition and regular inundation by the tides. Succulent leaves, stilt roots, pneumatophores, and viviparous germination are the key features of these plants. Heritiera fomes, Excoecaria agallocha, Nypa fruticans, Sonneratia apetala, Rhizophora spp., Ceriops decandra, Phoenix paludosa, and Acrostichum aureum are the common plants of the Sundarbans. Wetlands of Bangladesh also support a large number of plants and wild animals of the country. Nearly 50% (8 million ha) of the total land surface of the country including river, natural lakes, tanks, reservoir, mangrove forests, estuarine, and seasonally inundated floodplains are considered as wetland.
\nBangladesh is situated in the northeastern part of the South Asia region, lying between 20°25′ and 26°38′ north latitude and 88°01′ and 92°40′ east longitude. The country is bordered by India to the north, northeast, and west, Myanmar to the southeast, and the Bay of Bengal covering the southern part with a coastline of 710 km. The climate of Bangladesh is tropical, with maximum summer temperature rising between 32 and 38°C. Annual rainfall ranges between 200 and 400 cm. Biogeographically the country lies at the junction of the Indian and Malayan subregions of the Indo-Malayan Realm and is located very near to the western side of Sino-Japanese region.
\nProtected areas are “areas especially dedicated to the protection and maintenance of biological diversity and associated cultural resources, which are managed through legal or other effective means, designated or regulated and managed to achieve specific conservation objectives” [12, 13]. PAs have long been considered as the cornerstone of all national and regional conservation strategies. While it is often argued that they are the most effective and widespread measure for conserving forests and biodiversity [13, 14], the importance of complementary off-reserve management has also been acknowledged [15, 16, 17]. Globally, the number of PAs has increased significantly over the last few decades in recognition of their importance for conservation. At present, there are more than 100,000 protected area sites worldwide, covering nearly 12% of the world’s land surface [18, 19, 20]. Currently there are 34 forest PAs in Bangladesh which represent about 17.5% of the total forest land of the country and 1.8% of country’s total land [21].
\nBangladesh Forest Department shifts its paradigm of conventional forest management to community-led management for ensuring effective governance of natural resources. A total of 34 reserved forests have been declared as protected area where 23 PAs are managed through active participation of community people which is known as co-management. Community people come forward along with the forest department regarding conservation through ensuring sustainable natural resource management.
\nThe forestry sector of Bangladesh plays an important role in combating poverty for the people living in and around the forest. The history of forest management in Bangladesh is quite old and was shaped and influenced by colonial forest policy. The Forest Policy, 1979, was the first of its kind and was very much influenced by the colonial policy of forest management [22]. Over time this policy proved ineffective due to various socioeconomic factors such as population growth, poverty, overexploitation of resources, and top-down, centralized management approaches. It was felt by experts, communities, and policy makers that a new dimension to the existing forest policy was needed. The Forest Policy, 1994, specifically recognized the importance of peoples’ participation in forest management [23]. Another notable achievement of the 1994 policy was that it has succeeded in bringing tree plantation activities outside the forest area [24].
\nMost importantly, significant developments in Bangladesh forest legal and policy frameworks took place after the formulation of the 1994 policy [25]. Community participation in the forestry sector of Bangladesh has a long history that can be traced back as early as 1871, to teak plantations of Chittagong Hill Tracts managed by the tribal farmers. However, participation of community people in the forestry sector officially began in the 1980s. Donor-assisted community forestry project was the first attempt of its kind in the northwestern districts of Bangladesh. It gradually spread to other parts of the country through various projects and forms such as the Thana Afforestation and Nursery Development Project (TANDP), the Coastal Greenbelt Project (CGP), and the Forestry Sector Project (FSP). Despite the initial success in achieving physical targets, i.e., increase of plantation coverage, these projects failed to develop a mechanism to attract and engage local communities. They lacked institutional, personal, and community capacity building, legitimacy on usufruct rights, active community participation, and devolution of the decision-making power under the continued influence of “command and control” strategies. The introduction of co-management in the forest PAs is an effort to overcome these limitations to incorporate active community participation as a core aspect of PA governance [23]. The government of Bangladesh started introducing and implementing co-management in five forest PAs under a pilot project titled Nishorgo Support Project (hereafter referred to as NSP) for a period of 5 years (2004–2009) [26]. Many countries have already developed enabling legal and policy frameworks to support community rights and access and have thereby offered better incentives in the governance of the PAs and the resources sustained by them.
\nAs an effective tool, the protected areas are recognized internationally for the conservation of biodiversity. Currently the PAs of Bangladesh represent most of the ecosystems and thus include all habitats and species that are vital for conservation. The Bangladesh Forest Department under the Ministry of Environment, Forest and Climate Change manages a network of 17 national parks, 20 wildlife sanctuaries, 2 special biodiversity conservation areas, 1 marine protected area, 2 vulture safe zones, 2 botanical gardens, 2 safari parks, 10 eco-parks, and 1 aviary park. The total area under this protected area status covers 618253.49 ha of forest land and represents 4.19% area of the country [27]. The primary purpose of these sites is to conserve and protect habitat for wildlife, including migratory birds, species at risk, and other species of national interest.
\nAs a sample of protected areas of Bangladesh, we reviewed the floristic studies [28, 29, 30] conducted in three protected areas of Bangladesh with characteristic features. They are “Chunati Wildlife Sanctuary,” Dudhpukuria-Dhopachari Wildlife Sanctuary, and “Madhupur National Park.” We used the plant data collected during the field survey as secondary data for assessing the fodder yielding plants with due permission from the respective authors. The identified plants were then explored with their use and conservation status following the encyclopedia of flora and fauna of Bangladesh [31].
\nChunati Wildlife Sanctuary, established in 1986, is familiar as the habitat and breeding ground of the Asian elephant (Elephas maximus). It is one of the oldest PAs of Bangladesh rich with 691 plants from all habit forms [28]. In addition to the Asian elephant, Chunati harbors 26 species of amphibians, 54 reptiles, 252 birds, and 40 mammals [32]. Tables 2 and 3 provide a detailed account of the flora and fauna of the selected protected areas. Dudhpukuria-Dhopachari Wildlife Sanctuary is a comparatively new protected area that is declared in 2010. It covers an area of 4716 ha rich in both floral and faunal diversity. The wildlife sanctuary harbors 608 plant species and 385 wildlife [29]. However, the Asian elephant is also the flagship animal of this PA. Madhupur National Park, also one of the oldest protected areas, was declared in 1982. It is situated in the central region of Bangladesh covered with mainly deciduous Shorea robusta. It harbors 385 plant species from all habit forms and 192 wildlife including amphibians, birds, mammals, and reptiles (Tables 2 and 3).
\nWe identified the wild fodder yielding flora of different habit forms following the encyclopedia of flora and fauna of Bangladesh [31]. The review indicated that each of the protected areas harbors a substantial number of fodder yielding plants from different habit forms (Table 2).
\nA total of 112 tree species belonging to 71 genera and 32 families were found to yield part of it (i.e., leaves, branch, fruit, seed, flower, etc.) as fodder. A comparison number of species in the selected PAs indicated that CWS has 87 species, whereas DDWS and MNP showed 69 and 67 species, respectively (Table 4). Density of the fodder yielding tree species varied greatly with PAs. Ficus hispida was having the highest stem density in CWS, whereas in DDWS Grewia nervosa and Artocarpus chama were having the highest density. On the other hand, Mallotus philippensis and Protium serratum were the two mostly dense tree species in MNP. There were 15 fodder yielding exotic tree species in the three protected areas. The studies indicated that density of very few species was good (10 stems/ha); however most of them are having very poor density which apparently seems not indicative of a rich wildlife habitat.
\nHabit forms | \nNumber of species reported from the selected protected areas | \n|||||
---|---|---|---|---|---|---|
Chunati Wildlife Sanctuary | \nDudhpukuria-Dhopachari Wildlife Sanctuary | \nMadhupur National Park | \n||||
Total | \nFodder yielding | \nTotal | \nFodder yielding | \nTotal | \nFodder yielding | \n|
Trees | \n240 | \n81 | \n182 | \n61 | \n139 | \n70 | \n
Shrubs | \n102 | \n17 | \n125 | \n21 | \n48 | \n10 | \n
Herbs | \n211 | \n61 | \n200 | \n70 | \n136 | \n43 | \n
Climbers | \n106 | \n25 | \n71 | \n8 | \n46 | \n22 | \n
Ferns | \n19 | \n4 | \n17 | \n4 | \n9 | \n2 | \n
Epiphytes | \n7 | \n— | \n7 | \n— | \n7 | \n— | \n
Parasites | \n6 | \n— | \n6 | \n— | \n— | \n— | \n
Total | \n691 | \n188 | \n608 | \n164 | \n385 | \n147 | \n
Groups | \nNumber of species in the selected protected areas | \n||
---|---|---|---|
Chunati Wildlife Sanctuary | \nDudhpukuria-Dhopachari Wildlife Sanctuary | \nMadhupur National Park | \n|
Amphibians | \n26 | \n25 | \n17 | \n
Birds | \n252 | \n231 | \n120 | \n
Fish | \n10 | \n23 | \n— | \n
Mammals | \n40 | \n50 | \n27 | \n
Reptiles | \n54 | \n56 | \n28 | \n
SN | \nBotanical name | \nLocal name | \nFamily | \nConservation status | \nDensity in PAs (stem/ha) | \n||
---|---|---|---|---|---|---|---|
CWS | \nDDWS | \nMNP | \n|||||
1 | \nAcacia mangium Willd. | \nMangium | \nMimosaceae | \nLC | \n24.3 | \n0.4 | \n0.4 | \n
2 | \nAcronychia pedunculata (L.) Miq. | \nBonjamir, Jairgola | \nRutaceae | \nNE | \n0.2 | \n10.6 | \n\n |
3 | \nAegle marmelos (L.) Corr. | \nBel | \nRutaceae | \nLC | \n0.1 | \n0.4 | \n0.4 | \n
4 | \nAlangium chinense (Lour.) Harms | \nMarleza Gachh | \nAlangiaceae | \nNE (rare) | \n\n | 3.6 | \n\n |
5 | \nAlbizia chinensis (Osb.) Merr. | \nChakua Koroi | \nMimosaceae | \nLC | \n4 | \n4.8 | \n0.9 | \n
6 | \nAlbizia lebbeck (L.) Benth. & Hook | \nKala Koroi | \nMimosaceae | \nLC | \n\n | \n | 1.3 | \n
7 | \nAlbizia odoratissima (L. f.) Benth. | \nTetoyakoroi | \nMimosaceae | \nLC | \n0.4 | \n4.4 | \n\n |
8 | \nAnacardium occidentale L. | \nKajubadam | \nAnacardiaceae | \nLC | \n0.8 | \n\n | \n |
9 | \nAnnona squamosa L. | \nAta | \nAnnonaceae | \nLC | \n0.4 | \n\n | 0.4 | \n
10 | \nAntidesma acidum Retz. | \nElena | \nEuphorbiaceae | \nLC | \n0.8 | \n0.4 | \n\n |
11 | \nAntidesma acuminatum Wall. in Wight. | \nChokoi | \nEuphorbiaceae | \nLC | \n\n | \n | 0.4 | \n
12 | \nAntidesma bunius (L.) Spreng. | \nWishwar choa, Banshial Boka | \nEuphorbiaceae | \nLC | \n1.3 | \n0.4 | \n\n |
13 | \nAntidesma ghaesembilla Gaertn. | \nChokoi, Elena | \nEuphorbiaceae | \nLC | \n0.2 | \n\n | 2.6 | \n
14 | \nAphanamixis polystachya (Wall.) R.N. Parker. | \nPtiraj | \nMimosaceae | \nLC | \n0.8 | \n3.8 | \n0.4 | \n
15 | \nAporosa dioica (Roxb.) Mull.Arg. | \nCastoma | \nEuphorbiaceae | \nNE | \n3.8 | \n\n | \n |
16 | \nAporosa sp. | \nKharjon | \nEuphorbiaceae | \nNE | \n\n | \n | 23.3 | \n
17 | \nAreca catechu L. | \nSupari | \nArecaceae | \nLC | \n0.2 | \n\n | 0.4 | \n
18 | \nArtocarpus chama Buch.-Ham. ex Wall. | \nChapalish, Chambal | \nMoraceae | \nNE (rare) | \n3.9 | \n17.6 | \n0.4 | \n
19 | \nArtocarpus heterophyllus Lamk. | \nKanthal | \nMoraceae | \nNE (rare) | \n4.3 | \n0.2 | \n0.4 | \n
20 | \nArtocarpus lacucha Buch.-Ham. | \nBorta | \nMoraceae | \nLC | \n2.4 | \n4.4 | \n1.3 | \n
21 | \nAverrhoa bilimbi L. | \nBelombo | \nOxalidaceae | \nLC | \n0.2 | \n\n | \n |
22 | \nAverrhoa carambola L. | \nKamranga | \nOxalidaceae | \nLC | \n0.2 | \n\n | \n |
23 | \nBaccaurea ramiflora Lour. | \nLotkon | \nEuphorbiaceae | \nLC | \n0.1 | \n0.2 | \n\n |
24 | \nBorassus flabellifer L. | \nTal | \nArecaceae | \nLC | \n0.2 | \n\n | 0.4 | \n
25 | \nBridelia retusa (L.) A. Juss. | \nKata Kushui, Kata Koi | \nEuphorbiaceae | \nLC | \n\n | 0.6 | \n\n |
26 | \nBuchanania lancifolia Roxb. | \n\n | Anacardiaceae | \nNE (rare) | \n0.2 | \n\n | \n |
27 | \nCallicarpa arborea Roxb. | \nBormala, Khoja | \nVerbenaceae | \nLC | \n7.4 | \n6.8 | \n0.4 | \n
28 | \nCalophyllum polyanthum Wall. ex Choisy | \nChandua, Kamdeb | \nClusiaceae | \nNE (rare) | \n\n | 0.6 | \n\n |
29 | \nCassia fistula L. | \nSonalu | \nCaesalpiniaceae | \nLC | \n0.5 | \n1 | \n1.7 | \n
30 | \nCitrus reticulata Blanco | \nKomla | \nRutaceae | \nLC | \n0.1 | \n\n | 0.4 | \n
31 | \nClausena heptaphylla (Roxb.) Wight & Arn. ex Steud. | \nKaran phal, Panbahar, sada Moricha | \nRutaceae | \nLC | \n1.12 | \n0.2 | \n\n |
32 | \nCleistocalyx nervosum (DC.) Kosterm. var. paniala (Roxb.) J. Parn. & P. Chantaranothai | \n\n | Myrtaceae | \nLC | \n0.2 | \n\n | 2.2 | \n
33 | \nCocos nucifera L. | \nNarikel | \nArecaceae | \nLC | \n0.2 | \n0.6 | \n0.4 | \n
34 | \nCordia dichotoma Forst. f. | \nBolla gota, Bohal, Bhola | \nBoraginaceae | \nLC | \n\n | \n | 0.4 | \n
35 | \nCordia dichotoma Forst. f. | \nBohal | \nBoraginaceae | \nLC | \n0.3 | \n\n | 0.4 | \n
36 | \nCrateva magna (Lour.) DC. | \n\n | Capparaceae | \nLC | \n0.3 | \n\n | \n |
37 | \nCryptocarya amygdalina Nees. | \nOjha | \nLauraceae | \nNE (rare) | \n2.5 | \n3.4 | \n21.9 | \n
38 | \nDalbergia sissoo Roxb. | \nSissoo | \nFabaceae | \nLC | \n0.1 | \n\n | \n |
39 | \nDillenia indica L. | \nChalta | \nDilleniaceae | \nLC | \n0.2 | \n0.2 | \n\n |
40 | \nDillenia scabrella Roxb. ex Wall. | \nAjuli, Ajugi | \nDilleniaceae | \nLC | \n4.1 | \n5.8 | \n0.4 | \n
41 | \nDiospyros blancoi A. DC. | \nBilati gab | \nEbenaceae | \nLC | \n0.2 | \n\n | 0.4 | \n
42 | \nDiospyros malabarica (Desr.) Kostel. | \nDeshi gab | \nEbenaceae | \nLC | \n0.2 | \n1.4 | \n\n |
43 | \nElaeis guineensis Jacq. | \nPalm oil | \nArecaceae | \nNE | \n0.3 | \n\n | 0.4 | \n
44 | \nElaeocarpus floribundus Blume | \nTitpai | \nElaeocarpaceae | \nLC | \n0.2 | \n1.8 | \n1.3 | \n
45 | \nElaeocarpus tectorius (Lour.) Poir. | \nJalpai | \nElaeocarpaceae | \nLC | \n2.5 | \n2.2 | \n\n |
46 | \nFicus auriculata Lour. | \nLal Dumur | \nMoraceae | \nLC | \n1.6 | \n0.8 | \n\n |
47 | \nFicus benghalensis L. | \nBot | \nMoraceae | \nLC | \n0.6 | \n1.2 | \n1.7 | \n
48 | \nFicus hispida L. f. | \nDumur | \nMoraceae | \nLC | \n26.9 | \n4.6 | \n0.9 | \n
49 | \nFicus lanceolata Buch.-Ham. ex Roxb. | \n\n | Moraceae | \nV | \n0.3 | \n\n | \n |
50 | \nFicus racemosa L. | \nDumur, Jagyadumur | \nMoraceae | \nLC | \n0.3 | \n2.4 | \n0.9 | \n
51 | \nFicus religiosa L. | \nBot | \nMoraceae | \nLC | \n\n | \n | 0.4 | \n
52 | \nFicus rumphii Bl. | \nBot | \nMoraceae | \nLC | \n\n | \n | 0.4 | \n
53 | \nFicus semicordata Buch.-Ham. ex Smith | \nChokorgola | \nMoraceae | \nNE | \n0.9 | \n0.8 | \n\n |
54 | \nFicus tinctoria G. Forst. subsp. gibbosa (Blume) Corner | \n\n | Moraceae | \nNE (rare) | \n0.1 | \n\n | \n |
55 | \nFicus variegata Blume | \n\n | Moraceae | \nNE | \n0.4 | \n5.2 | \n\n |
56 | \nFicus virens Ait. | \nPakur, Pakar, Paikur | \nMoraceae | \nLC | \n0.1 | \n\n | 0.9 | \n
57 | \nFirmiana colorata (Roxb.) R. Br. | \nUdal | \nSterculiaceae | \nLC | \n0.2 | \n\n | \n |
58 | \nFlacourtia jangomas (Lour.) Raeusch. | \nPainnagola | \nFlacourtiaceae | \nLC | \n1.2 | \n0.2 | \n0.9 | \n
59 | \nGarcinia cowa Roxb. ex DC. | \nCao | \nClusiaceae | \nNE (rare) | \n9.3 | \n5.2 | \n2.2 | \n
60 | \nGarcinia lanceaefolia Roxb. | \n\n | Clusiaceae | \nNE | \n0.2 | \n\n | \n |
61 | \nGarcinia xanthochymus Hook. f. ex T. Anders. | \nTamal, Dephal | \nClusiaceae | \nLC | \n\n | 1 | \n\n |
62 | \nGaruga pinnata Roxb. | \nBhadi, Silbhadi, Jeolbhadi | \nBurseraceae | \nLC | \n0.2 | \n7 | \n3.9 | \n
63 | \nGrewia nervosa (Lour.) Panigr. | \nDatoi | \nTiliaceae | \nLC | \n8.5 | \n19.2 | \n22.4 | \n
64 | \nGrewia sapida Roxb. ex DC. | \nNaricha | \nTiliaceae | \nLC | \n0.2 | \n\n | \n |
65 | \nGrewia tiliifolia Vahl. | \nPholsa, Dhomoni | \nTiliaceae | \nLC | \n\n | 0.6 | \n\n |
66 | \nGrewia serrulata DC. | \nNaricha | \nTiliaceae | \nLC | \n\n | \n | 0.4 | \n
67 | \nHevea brasiliensis (Willd. ex A. Juss.) Mull.Arg. | \nRubber | \nEuphorbiaceae | \nLC | \n\n | \n | 0.4 | \n
68 | \nHydnocarpus laurifolius (Dennst.) Sleum. | \nHiddigach | \nFlacourtiaceae | \nNE (rare) | \n1.4 | \n5.2 | \n\n |
69 | \nLannea coromandelica (Houtt.) Merr. | \nJialbhadi | \nAnacardiaceae | \nLC | \n4 | \n0.2 | \n3.9 | \n
70 | \nLepisanthes rubiginosa (Roxb.) Leenh. | \n\n | Sapindaceae | \nLC | \n0.2 | \n0.2 | \n\n |
71 | \nLepisanthes senegalensis (Poir.) Leenh. | \nGotaharina | \nSapindaceae | \nLC | \n0.2 | \n0.2 | \n\n |
72 | \nLitchi chinensis Sonn. | \nLitchu, Lychee | \nSapindaceae | \nLC | \n1.0 | \n\n | 0.4 | \n
73 | \nMaesa indica (Roxb.) A. DC. | \nMaesa, Moricha, Romjani | \nMyrsinaceae | \nCD | \n0.1 | \n2.2 | \n\n |
74 | \nMallotus philippensis (Lamk.) Mull.Arg. | \nSinduri | \nEuphorbiaceae | \nCD | \n0.6 | \n0.2 | \n60. 8 | \n
75 | \nMangifera indica L. | \nAm | \nAnacardiaceae | \nLC | \n2.9 | \n0.4 | \n0.9 | \n
76 | \nMangifera sylvatica Roxb. | \nUriam | \nAnacardiaceae | \nV | \n0.1 | \n0.2 | \n\n |
77 | \nManilkara zapota (L.) P. van Royen | \nSofeda | \nSapotaceae | \nLC | \n\n | \n | 0.4 | \n
78 | \nMiliusa velutina (Dunal) Hook. f. | \nGandhi gajari | \nAnnonaceae | \nLC | \n\n | \n | 3.0 | \n
79 | \nMoringa oleifera Lamk. | \nSajna | \nMoringaceae | \nNE | \n\n | \n | 0.9 | \n
80 | \nPeltophorum pterocarpum (DC.) K. Heyne | \nRadhachura, Halud Krisnachura | \nCaesalpiniaceae | \nLC | \n0.4 | \n\n | 0.4 | \n
81 | \nPhoebe lanceolata (Ness) Ness | \nChaongri, Dulia | \nLauraceae | \nNE | \n\n | 0.2 | \n\n |
82 | \nPhoenix acaulis Roxb. | \nBon Khejur, Khudi khejur | \nArecaceae | \nV | \n\n | \n | 0.4 | \n
83 | \nPhoenix sylvestris Roxb. | \nKhejur | \nArecaceae | \nLC | \n0.2 | \n0.4 | \n0.4 | \n
84 | \nPhyllanthus emblica L. | \nAmloki | \nEuphorbiaceae | \nLC | \n2.4 | \n2 | \n0.9 | \n
85 | \nProtium serratum (Wall. ex Colebr.) Engl. | \nGotgutia | \nBurseraceae | \nLC | \n2.1 | \n12.2 | \n32.8 | \n
86 | \nPsidium guajava L. | \nPayara | \nMyrtaceae | \nLC | \n4.2 | \n0.2 | \n0.4 | \n
87 | \nSamanea saman (Jacq.) Merr. | \nRaintree | \nMimosaceae | \nLC | \n0.4 | \n\n | 0.9 | \n
88 | \nSapium baccatum Roxb. | \nCham phata | \nEuphorbiaceae | \nLC | \n0.4 | \n3 | \n\n |
89 | \nSchleichera oleosa (Lour.) Oken. | \nJoyna, Kusum | \nSapindaceae | \nNE | \n\n | \n | 14.2 | \n
90 | \nSemecarpus anacardium L.f. | \nBheula, Bhela | \nAnacardiaceae | \nNE | \n\n | \n | 11.2 | \n
91 | \nSenna siamea (Lamk.) Irwin & Barneby | \nMinjiri | \nCaesalpiniaceae | \nLC | \n3.6 | \n1.8 | \n0.4 | \n
92 | \nSpondias pinnata (L.f.) Kurz | \nBon-Amra, Piala | \nAnacardiaceae | \nLC | \n\n | 3.4 | \n0.4 | \n
93 | \nSterculia hamiltonii (O. Kuntze) Adelb. | \n\n | Sterculiaceae | \nLC | \n\n | 0.2 | \n\n |
94 | \nStreblus asper Lour. | \nSheora, Harba | \nMoraceae | \nLC | \n1.5 | \n2 | \n0.9 | \n
95 | \nSyzygium balsameum (Wight) Walp. | \nButi Jam | \nMyrtaceae | \nLC | \n\n | 1 | \n\n |
96 | \nSyzygium claviflorum (Roxb.) A. M. Cowan & J. M. Cowan | \n\n | Myrtaceae | \nLC | \n3.8 | \n0.4 | \n\n |
97 | \nSyzygium cumini (L.) Skeels | \nKalojam | \nMyrtaceae | \nLC | \n1.3 | \n0.8 | \n0.9 | \n
98 | \nSyzygium cymosum DC. | \nKhudi Jam | \nMyrtaceae | \nNE | \n\n | 0.2 | \n\n |
99 | \nSyzygium firmum Thw. | \nDhaki jam | \nMyrtaceae | \nLC | \n7.5 | \n1.8 | \n0.4 | \n
100 | \nSyzygium fruticosum (Wall.) Masamune | \nPutijam | \nMyrtaceae | \nLC | \n13.0 | \n1 | \n6.5 | \n
101 | \nSyzygium jambos (L.) Alston | \nGulapjam | \nMyrtaceae | \nLC | \n0.2 | \n\n | \n |
102 | \nSyzygium praecox (Roxb.) Rathakr. & N. C. Nair | \n\n | Myrtaceae | \nNE | \n0.2 | \n\n | \n |
103 | \nSyzygium tetragonum Wall. ex Kurz. | \nPholda jam, Lal Pholda | \nMyrtaceae | \nNE | \n\n | 3 | \n\n |
104 | \nTamarindus indica L. | \nTentul | \nCaesalpiniaceae | \nLC | \n0.1 | \n0.6 | \n0.9 | \n
105 | \nTerminalia bellirica (Gaertn.) Roxb. | \nBohera | \nCombretaceae | \nLC | \n7.5 | \n10.6 | \n29.3 | \n
106 | \nTerminalia catappa L. | \nKatbadam | \nCombretaceae | \nLC | \n0.2 | \n\n | \n |
107 | \nTerminalia chebula Retz. | \nHaritaki | \nCombretaceae | \nV | \n0.8 | \n0.6 | \n4.7 | \n
108 | \nTetrameles nudiflora R. Br. | \nChandul, Maina Kat | \nDatiscaceae | \nNE | \n0.1 | \n1.8 | \n\n |
109 | \nTrema orientalis (L.) Blume | \nJiban, Naricha | \nUlmaceae | \nLC | \n5.1 | \n0.2 | \n0.4 | \n
110 | \nVitex glabrata R.Br. | \nGoda arsol, Hakuni gach | \nVerbenaceae | \nLC | \n1.0 | \n1.4 | \n0.9 | \n
111 | \nVitex peduncularis Wall. ex Schauer | \nGoda | \nVerbenaceae | \nNE (rare) | \n3.3 | \n11 | \n0.9 | \n
112 | \nZiziphus mauritiana Lamk. | \nBoroi | \nRhamnaceae | \nLC | \n0.2 | \n\n | 0.4 | \n
List of fodder yielding trees in three selected protected areas [here, LC, least concern; NE, not evaluated; NE (rare), not evaluated but seems to be rare].
There were 27 fodder yielding shrubby species recorded from the selected three protected areas. These species taxonomically belong to 23 genera and 15 families (Table 5). Both CWS and DDWS were represented by 17 shrubby fodder yielding species, whereas MNP showed 14 species indicating its comparative inferiority of supporting wildlife. However, Cajanus cajan and Manihot esculenta were the two exotic fodder species recorded from the cultivation sites of MNP and CWS.
\nSN | \nBotanical name | \nLocal name | \nFamily | \nConservation status | \nOccurrence in selected PAs | \n||
---|---|---|---|---|---|---|---|
CWS | \nDDWS | \nMNP | \n|||||
1 | \nBambusa tulda Roxb. | \nMitinga, Mitinga, Mirtinga, Taralla, Tolla bansh | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
2 | \nBambusa vulgaris Schrad. ex Wendl. | \nBaijja, Baria, Jowa Bansh, Bangla Bans, Ora Bansh | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
3 | \nBauhinia acuminata L. | \n\n | Caesalpiniaceae | \nLC | \n✓ | \n\n | \n |
4 | \nBridelia stipularis (L.) Blume | \nSitki (climbing) | \nEuphorbiaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
5 | \nCaesalpinia hymenocarpa (Prain) Hattink. | \n\n | Caesalpiniaceae | \nNE (rare) | \n\n | ✓ | \n\n |
6 | \nCajanus cajan (L.) Millsp. | \nArhor, Sarata alu, Sortai alu | \nFabaceae | \nLC | \n✓ | \n\n | ✓ | \n
7 | \nCapparis zeylanica L. | \n\n | Capparaceae | \nLC | \n✓ | \n✓ | \n\n |
8 | \nCitrus aurantifolia (Christm. & Panzer) Swingle | \nLebu | \nRutaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
9 | \nClausena suffruticosa (Roxb.) Wight & Arn. | \nSadamoricha | \nRutaceae | \nLC | \n✓ | \n\n | \n |
10 | \nClerodendrum serratum (L.) Moon | \n\n | Verbenaceae | \nNE | \n✓ | \n\n | \n |
11 | \nCrotalaria spectabilis Roth | \nPipli-jhunjan | \nFabaceae | \nLC | \n\n | ✓ | \n\n |
12 | \nDendrocalamus longispathus (Kurz) Kurz | \nOra | \nPoaceae | \nNE | \n\n | ✓ | \n\n |
13 | \nGrewia asiatica L. | \nPholsa | \nTiliaceae | \nNE (rare) | \n\n | ✓ | \n✓ | \n
14 | \nGrewia serrulata DC. | \nPanisara, Pichandi, Khulla damor | \nTiliaceae | \nLC | \n\n | ✓ | \n✓ | \n
15 | \nHelicia erratica Hook. f. | \n\n | Proteaceae | \nNT | \n✓ | \n\n | \n |
16 | \nMaclura cochinchinensis (Lour.) Corner | \n\n | Moraceae | \nLC | \n\n | ✓ | \n\n |
17 | \nManihot esculenta Crantz | \nCassava, Gach alu | \nEuphorbiaceae | \nLC | \n\n | \n | ✓ | \n
18 | \nMelocanna baccifera (Roxb.) Kurz | \nMuli | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
19 | \nMurraya koenigii (L.) Spreng. | \n\n | Rutaceae | \nLC | \n✓ | \n\n | \n |
20 | \nPhlogacanthus thyrsiformis Roxb ex D. J. Mabberley | \n\n | Acanthaceae | \nNE | \n\n | \n | ✓ | \n
21 | \nPremna esculenta Roxb. | \nLalana | \nVerbenaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
22 | \nPunica granatum L. | \nDalim | \nPunicaceae | \nLC | \n\n | \n | ✓ | \n
23 | \nSarcochlamys pulcherrima Gaudich. | \nJangallya shak, Maricha | \nUrticaceae | \nNE | \n✓ | \n✓ | \n\n |
24 | \nSolanum melongena L. | \nBegun | \nSolanaceae | \nLC | \n✓ | \n\n | ✓ | \n
25 | \nSolanum torvum Sw. | \nTit begun, Gota begun | \nSolanaceae | \nLC | \n✓ | \n✓ | \n\n |
26 | \nZiziphus oenoplia (L.) Mill. | \nBonboroi, Toktoki kanta, Tokni boroi | \nRhamnaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
27 | \nZiziphus rugosa Lamk. | \nJangli Boroi, Anoi, Anoi gota, Anari gota | \nRhamnaceae | \nNE | \n\n | ✓ | \n✓ | \n
List of fodder yielding shrubs reported from the selected protected areas [here, LC, least concern; NE, not evaluated; NE (rare), not evaluated but seems to be rare].
The review revealed a total of a higher number of fodder yielding herbs occurring in the protected areas. One hundred twenty-one herbaceous species belonging to 82 genera and 29 families were recorded from the protected areas (Table 6). DDWS was represented with the highest number of herb species (70 species) which was followed by CWS and MNP with 60 and 39 herb species, respectively. A substantial number of (15 species) herbs that are reported growing in and around the protected areas were introduced in Bangladesh at different times, and most of these were found to be cultivated in the adjacent forest areas of the protected areas. Wildlife takes advantages of cultivation by raiding them for food especially during the cultivation season. The conservation status of three fodder yielding herbs was vulnerable, i.e., Colocasia oresbia, Homalomena coerulescens, and Polygala furcata.
\nSN | \nBotanical name | \nLocal name | \nFamily | \nConservation status | \nOccurrence in selected PAs | \n||
---|---|---|---|---|---|---|---|
CWS | \nDDWS | \nMNP | \n|||||
1 | \nAcroceras tonkinense (Balansa) C.E. Hubb. ex Bor | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
2 | \nActinoscirpus grossus (L.f.) Goetgh & D. A. Simpson | \nKasuru, Kasari, Kesar | \nCyperaceae | \nLC | \n\n | ✓ | \n\n |
3 | \nAllium cepa L. | \nPiyaj | \nLiliaceae | \nLC | \n✓ | \n\n | \n |
4 | \nAllium sativum L. | \nRashun | \nLiliaceae | \nLC | \n✓ | \n\n | \n |
5 | \nAlocasia macrorrhizos (L.) G. Don | \nMankatchu | \nAraceae | \nLC | \n\n | \n | ✓ | \n
6 | \nAlternanthera philoxeroides (Mart.) Griseb. | \nHelencha, Malancha shak | \nAmaranthaceae | \nLC | \n\n | \n | ✓ | \n
7 | \nAmaranthus spinosus L. | \nKantashakh, Kata Notay, Khoira kanta | \nAmaranthaceae | \nLC | \n✓ | \n\n | ✓ | \n
8 | \nAmaranthus tricolor L. | \nLalshakh, Danga, Data shak | \nAmaranthaceae | \nLC | \n✓ | \n\n | ✓ | \n
9 | \nAmischophacelus axillaris (L.) Rolla Rao & Kamm. | \nBaghanulla | \nCommelinaceae | \nLC | \n\n | ✓ | \n\n |
10 | \nAmorphophallus bulbifer (Roxb.) Blume | \nAmla-bela, Jongle Ol. | \nAraceae | \nLC | \n\n | ✓ | \n✓ | \n
11 | \nAmorphophallus paeoniifolius (Dennst.) Nicolson | \nBag katchu, Batema katchu | \nAraceae | \nLC | \n\n | \n | ✓ | \n
12 | \nAnanas comosus (L.) Merr. | \nAnarosh | \nBromeliaceae | \nLC | \n✓ | \n\n | ✓ | \n
13 | \nAponogeton echinatus Roxb. | \nGhechu | \nAponogetonaceae | \nCD | \n\n | ✓ | \n\n |
14 | \nAponogeton natans (L.) Engl. & Krause | \n\n | Aponogetonaceae | \nNT | \n\n | ✓ | \n\n |
15 | \nArundo donax L. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
16 | \nAxonopus compressus (Sw.) P. Beauv. | \nGhora dubo Har, Farak pata | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
17 | \nBothriochloa bladhii (Retz.) S. T. Blake | \nGandha Gourana | \nPoaceae | \nLC | \n\n | ✓ | \n\n |
18 | \nBrachiaria decumbens Stapf | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
19 | \nBrachiaria distachya (L.) Stapf | \nCorighas | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
20 | \nBrachiaria kurzii (Hook. f.) A. Camus | \n\n | Poaceae | \nLC | \n\n | \n | ✓ | \n
21 | \nBrachiaria reptans (L.) Gard. & Hubb. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
22 | \nBrassica oleracea L. var. botrytis L. | \nPhulkopi | \nBrassicaceae | \nLC | \n✓ | \n\n | \n |
23 | \nBryophyllum pinnatum (Lamk.) Oken | \nPathorkutchi, Pathorchura | \nCrassulaceae | \nLC | \n✓ | \n\n | ✓ | \n
24 | \nCapillipedium assimile (Steud.) A Camus | \n\n | Poaceae | \nDD | \n\n | ✓ | \n\n |
25 | \nCapsicum annuum L. | \nMorich | \nSolanaceae | \nLC | \n\n | \n | ✓ | \n
26 | \nCapsicum frutescens L. | \nMorich | \nSolanaceae | \nLC | \n✓ | \n\n | \n |
27 | \nCarica papaya L. | \nPapaya, Pape | \nCaricaceae | \nLC | \n✓ | \n\n | ✓ | \n
28 | \nCentella asiatica (L.) Urban | \nThankuni | \nApiaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
29 | \nChenopodium album L. | \nBatua Shakh | \nChenopodiaceae | \nLC | \n✓ | \n\n | \n |
30 | \nChrysopogon aciculatus (Retz.) Trin | \nLengra, Premkanta | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
31 | \nColocasia esculenta (L.) Schott | \nKatchu | \nAraceae | \nLC | \n✓ | \n✓ | \n✓ | \n
32 | \nColocasia fallax Schott | \n\n | Araceae | \nLC | \n\n | ✓ | \n\n |
33 | \nColocasia oresbia A. Hay | \nSadakachu | \nAraceae | \nV | \n\n | ✓ | \n\n |
34 | \nCommelina benghalensis L. | \nDholpata, Kanchira | \nCommelinaceae | \nLC | \n\n | ✓ | \n✓ | \n
35 | \nCommelina sikkimensis C.B. Clarke | \nBatbaithia Shag | \nCommelinaceae | \nCD | \n\n | ✓ | \n\n |
36 | \nCorchorus capsularis L. | \nPat shakh | \nTiliaceae | \nLC | \n\n | \n | ✓ | \n
37 | \nCurcuma longa L. | \nHalud | \nZingiberaceae | \nLC | \n✓ | \n\n | ✓ | \n
38 | \nCyanotis cristata (L.) D. Don | \n\n | Commelinaceae | \nLC | \n\n | ✓ | \n\n |
39 | \nCymbopogon citratus (DC) Stapf | \nDhan Sabarang, Lemon Ghas | \nPoaceae | \nCD | \n\n | ✓ | \n\n |
40 | \nCynodon arcuatus J. S. Presl ex C. B. Presl | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
41 | \nCynodon dactylon (L.) Pers. | \nDurba grass | \nPoaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
42 | \nCyperus corymbosus Rottb. | \nGola Methi | \nCyperaceae | \nNE | \n\n | ✓ | \n\n |
43 | \nCyperus cyperoides (L.) O. Ktze. | \nKucha, Kusha | \nCyperaceae | \nLC | \n\n | ✓ | \n\n |
44 | \nCyperus difformis L. | \nBehua | \nCyperaceae | \nLC | \n\n | ✓ | \n✓ | \n
45 | \nCyperus digitatus Roxb. | \nBehua | \nCyperaceae | \nLC | \n\n | ✓ | \n\n |
46 | \nCyperus distans L. f. | \nPani Malanga | \nCyperaceae | \nLC | \n\n | ✓ | \n\n |
47 | \nCyperus laxus Lamk var. laxus | \n\n | Cyperaceae | \nLC | \n\n | ✓ | \n✓ | \n
48 | \nCyperus rotundus L. | \nMutha | \nCyperaceae | \nLC | \n\n | ✓ | \n\n |
49 | \nCyperus tuberosus Rottb. | \n\n | Cyperaceae | \nLC | \n✓ | \n✓ | \n\n |
50 | \nCyrtococcum oxyphyllum (Steud.) Stapf | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
51 | \nCyrtococcum patens (L.) A. Camus | \n\n | Poaceae | \nLC | \n✓ | \n✓ | \n\n |
52 | \nDactyloctenium aegyptium (L.) P. Beauv. | \nMakra | \nPoaceae | \nLC | \n\n | ✓ | \n\n |
53 | \nDesmostachya bipinnata (L.) Stapf | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
54 | \nDichanthium caricosum (L.) A. Camus | \nDetara | \nPoaceae | \nLC | \n\n | ✓ | \n\n |
55 | \nDigitaria bicornis (Lamk.) Roem. & Schult. ex Loud | \n\n | Poaceae | \nNE | \n\n | \n | ✓ | \n
56 | \nDigitaria sanguinalis (L.) Scop. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
57 | \nEchinochloa crus-galli (L.) P. Beauv. | \nBara Shama-ghas | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
58 | \nEichhornia crassipes (Mart.) Solms | \n\n | Pontederiaceae | \nLC | \n✓ | \n\n | \n |
59 | \nEleusine indica (L.) Gaertn. | \nMalankuri | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
60 | \nEragrostis ciliaris (L.) R. Br. | \n\n | Poaceae | \nLC | \n✓ | \n\n | ✓ | \n
61 | \nEragrostis lehmanniana Nees | \n\n | Poaceae | \nNE | \n\n | ✓ | \n\n |
62 | \nEragrostis tenella (L.) P. Beauv. ex Roem. & Schult. | \nKoni Ghas | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
63 | \nEragrostis unioloides (Retz.) Nees ex Steud. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
64 | \nEriochloa procera (Retz.) C. E. Hubb. | \n\n | Poaceae | \nLC | \n✓ | \n\n | ✓ | \n
65 | \nEuphorbia heterophylla L. | \n\n | Euphorbiaceae | \nNE (rare) | \n\n | ✓ | \n\n |
66 | \nFuirena umbellata Rottb. | \n\n | Cyperaceae | \nLC | \n\n | ✓ | \n\n |
67 | \nHomalomena coerulescens Jungh. | \n\n | Araceae | \nV | \n\n | ✓ | \n\n |
68 | \nHydrolea zeylanica (L.) Vahl | \n\n | Hydrophyllaceae | \nLC | \n✓ | \n\n | \n |
69 | \nHymenachne pseudointerrupta C. Muell. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
70 | \nImperata cylindrica (L.) P. Beauv. | \nChhan, Chau, Kash | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
71 | \nJuncus prismatocarpus R.Br. | \n\n | Juncaceae | \nLC | \n\n | ✓ | \n✓ | \n
72 | \nKyllinga brevifolia Rottb. | \n\n | Cyperaceae | \nLC | \n\n | ✓ | \n\n |
73 | \nKyllinga bulbosa Beauv. | \n\n | Cyperaceae | \nLC | \n\n | ✓ | \n\n |
74 | \nKyllinga nemoralis (J. R. Forst. & G. Forst.) Dandy ex Hutchins. & Dalziel | \n\n | Cyperaceae | \nLC | \n✓ | \n✓ | \n\n |
75 | \nLasia spinosa (L.) Thw. | \n\n | Araceae | \nLC | \n✓ | \n\n | \n |
76 | \nLemna perpusilla Torrey | \n\n | Lemnaceae | \nLC | \n\n | ✓ | \n\n |
77 | \nLeucas indica (L.) R.Br. ex Vatke | \nDandakolas, Haldusha, Sweetadrone | \nLamiaceae | \nLC | \n\n | ✓ | \n\n |
78 | \nLophatherum gracile Brongn. | \n\n | Poaceae | \nLC | \n\n | \n | ✓ | \n
79 | \nLycopersicon esculentum Mill. | \nTomato | \nSolanaceae | \nLC | \n✓ | \n\n | ✓ | \n
80 | \nMollugo pentaphylla L. | \n\n | Molluginaceae | \nLC | \n\n | \n | ✓ | \n
81 | \nMonochoria hastata (L.) Solms | \nBaranukha | \nPontederiaceae | \nLC | \n✓ | \n✓ | \n\n |
82 | \nMonochoria vaginalis (Burm. f.) Presl | \nNukha, Sarkachu | \nPontederiaceae | \nLC | \n✓ | \n✓ | \n\n |
83 | \nMusa ornata Roxb. | \nRamkola | \nMusaceae | \nCD | \n✓ | \n✓ | \n\n |
84 | \nMusa paradisiaca L. | \nChampa kola | \nMusaceae | \nLC | \n✓ | \n\n | ✓ | \n
85 | \nOplismenus burmannii (Retz.) P. Beauv. | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
86 | \nOryza sativa L. | \nDhan | \nPoaceae | \nCD | \n✓ | \n\n | ✓ | \n
87 | \nPanicum maximum Jacq. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
88 | \nPanicum paludosum Roxb. | \nBarti, Borali, Kalash Nar | \nPoaceae | \nLC | \n\n | ✓ | \n\n |
89 | \nPanicum repens L. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
90 | \nPaspalidium flavidum (Retz.) A. Camus | \nBolai Mandi, Karin Ghas | \nPoaceae | \nLC | \n\n | ✓ | \n\n |
91 | \nPaspalum conjugatum Bergius | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
92 | \nPaspalum longifolium Roxb. | \n\n | Poaceae | \nNE | \n\n | ✓ | \n\n |
93 | \nPaspalum orbiculare G. Forst. | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
94 | \nPaspalum scrobiculatum L. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
95 | \nPhragmites karka (Retz.) Trin. ex Steud. | \nDharma, Nalkhagra | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
96 | \nPhysalis angulata L. | \nFotka | \nSolanaceae | \nLC | \n\n | ✓ | \n\n |
97 | \nPolygala furcata Royle | \n\n | Polygalaceae | \nV | \n\n | ✓ | \n\n |
98 | \nPueraria peduncularis (Grah. ex Benth.) Benth. | \n\n | Fabaceae | \nLC | \n\n | ✓ | \n\n |
99 | \nRaphanus sativus L. | \nMula | \nBrassicaceae | \nLC | \n✓ | \n\n | \n |
100 | \nRhynchospora corymbosa (L.) Britton | \n\n | Cyperaceae | \nLC | \n✓ | \n✓ | \n\n |
101 | \nSaccharum officinarum L. | \nAkh | \nPoaceae | \nCD | \n✓ | \n\n | \n |
102 | \nSaccharum ravennae L. | \nEkor | \nPoaceae | \nDD | \n✓ | \n\n | \n |
103 | \nSaccharum spontaneum L. | \nKash, Kaichcha, Kagara | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
104 | \nSacciolepis indica (L.) A. Chase | \n\n | Poaceae | \nLC | \n\n | ✓ | \n\n |
105 | \nSagittaria sagittifolia L. | \nMuyamuya, Chhotokut | \nAlismataceae | \nLC | \n\n | ✓ | \n\n |
106 | \nSenna hirsuta (L.) Irwin & Barneby | \n\n | Caesalpiniaceae | \nNE | \n\n | ✓ | \n\n |
107 | \nSesbania bispinosa (Jacq.) Wight. | \n\n | Fabaceae | \nLC | \n✓ | \n\n | \n |
108 | \nSetaria sphacelata (Schum.) Stapf. & C.E. Hubb. ex M. B. Moss | \n\n | Poaceae | \nCD | \n\n | ✓ | \n\n |
109 | \nSetaria verticillata (L.) P. Beauv. | \n\n | Poaceae | \nLC | \n✓ | \n\n | \n |
110 | \nSolanum americanum Mill. | \nTit-begun | \nSolanaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
111 | \nSolanum tuberosum L. | \nGolalu | \nSolanaceae | \nLC | \n✓ | \n\n | \n |
112 | \nSorghum bicolor (L.) Moench | \nJowar | \nPoaceae | \nCD | \n\n | \n | ✓ | \n
113 | \nThysanolaena maxima (Roxb.) Kuntze | \nJahruful | \nPoaceae | \nLC | \n✓ | \n✓ | \n\n |
114 | \nTyphonium trilobatum (L.) Schott | \nGhetkul | \nAraceae | \nLC | \n✓ | \n\n | ✓ | \n
115 | \nVernonia cinerea (L.) Less. | \nShial lata, Dankuni, Kuksim | \nAsteraceae | \nLC | \n✓ | \n✓ | \n✓ | \n
116 | \nXanthium indicum Koen. ex Roxb. | \nKhagra, Ban-okra | \nAsteraceae | \nLC | \n\n | \n | ✓ | \n
117 | \nXanthosoma sagittifolium (L.) Schott | \nPanchamukhi katchu | \nAraceae | \nLC | \n\n | \n | ✓ | \n
118 | \nXanthosoma violaceum Scott. | \nDudh katchu, Dastur | \nAraceae | \nLC | \n\n | \n | ✓ | \n
119 | \nZea mays L. | \nVuttra | \nPoaceae | \nCD | \n✓ | \n\n | \n |
120 | \nZingiber capitatum Roxb. | \nJongli Ada | \nZingiberaceae | \nNE (rare) | \n\n | ✓ | \n\n |
121 | \nZingiber officinale Rosc. | \nAda | \nZingiberaceae | \nLC | \n✓ | \n\n | ✓ | \n
List of fodder yielding herbs recorded from the selected protected areas [here, DD, data deficient; LC, least concern; NE, not evaluated; NE (rare), not evaluated but seems to be rare; V, vulnerable].
Climbers growing on bushes, trees, and forest floor are important fodder. The leaves and young shoots of the climbers and lianas are mainly eaten by wildlife as food. There were 39 climber plants belonging to 28 genera and 14 families reported as fodder species from the three protected areas. However, review of other reports indicates that the fodder yielding climber composition is almost similar to other forests of southeastern and central regions of Bangladesh. Besides the climber, there were seven fodder yielding pteridophytic species which belong to different genera of seven families. A total of five exotic climbers were recorded to be cultivated by the local people inside the boundary of the protected areas which sometimes raided by wildlife, i.e., monkey, squirrel, etc. The conservation status indicated the presence of one vulnerable plant named Calamus latifolius, the fruit of which is eaten by different birds and wildlife as food (Table 7).
\nSN | \nBotanical name | \nLocal name | \nFamily | \nConservation status | \nOccurrence in the PAs | \n||
---|---|---|---|---|---|---|---|
CWS | \nDDWS | \nMNP | \n|||||
1 | \nAcacia pennata (L.) Willd. | \nTeorakanta | \nMimosaceae | \nNE (rare) | \n\n | \n | ✓ | \n
2 | \nAmpelocissus barbata (Wallich) Planch. | \n\n | Vitaceae | \nCD | \n\n | \n | ✓ | \n
3 | \nAmpelocissus latifolia (Roxb.) Planch. | \n\n | Vitaceae | \nNE | \n\n | \n | ✓ | \n
4 | \nBasella rubra L. | \nPoi shak | \nBasellaceae | \nLC | \n\n | \n | ✓ | \n
5 | \nBenincasa hispida (Thunb.) Cogn. | \nChalkumra | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
6 | \nCaesalpinia digyna Rottler | \nKotchoi Kanta, Umulkuchi | \nCaesalpiniaceae | \nLC | \n✓ | \n✓ | \n\n |
7 | \nCalamus latifolius Roxb. | \nBudum bet, Korak bet | \nArecaceae | \nV | \n✓ | \n✓ | \n\n |
8 | \nCalamus tenuis Roxb. | \nChiringbet, Sanchi Bet, Bandari Bet, Jali bet | \nArecaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
9 | \nCissus elongata Roxb. | \n\n | Vitaceae | \nLC | \n\n | \n | ✓ | \n
10 | \nCitrullus lanatus (Thunb.) Matsumura & Nakai | \nTormuj | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
11 | \nCoccinia grandis (L.) Voigt | \nKawa jangi, Telakucha | \nCucurbitaceae | \nLC | \n\n | \n | ✓ | \n
12 | \nCoccinia grandis (L.) Voigt | \nTelakucha | \nCucurbitaceae | \nLC | \n✓ | \n✓ | \n\n |
13 | \nCucumis melo L. | \nKhira | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
14 | \nCucumis sativus L. | \nKhira, Futi | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
15 | \nCucurbita maxima Duch. ex Lamk. | \nMistikumra | \nCucurbitaceae | \nLC | \n✓ | \n\n | ✓ | \n
16 | \nDalbergia pinnata (Lour.) Prain | \nLalong-chhali, Keti | \nFabaceae | \nLC | \n\n | ✓ | \n\n |
17 | \nDioscorea belophylla (Prain) Voigt ex Haines | \nDudh alu | \nDioscoreaceae | \nLC | \n\n | \n | ✓ | \n
18 | \nDioscorea bulbifera L. | \nPagla Alu | \nDioscoreaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
19 | \nDioscorea esculenta (Lour.) Burkill | \nMaitta Alu | \nDioscoreaceae | \nNE | \n\n | \n | ✓ | \n
20 | \nDioscorea hamiltonii Hook. f. | \nThakan Budo | \nDioscoreaceae | \nNT | \n\n | ✓ | \n\n |
21 | \nDioscorea pentaphylla L. | \nAlu lata | \nDioscoreaceae | \nLC | \n✓ | \n✓ | \n✓ | \n
22 | \nIpomoea aquatica Forssk. | \nKalmi Shakh | \nConvolvulaceae | \nLC | \n✓ | \n\n | \n |
23 | \nIpomoea batatas (L.) Lamk. | \nMistialu | \nConvolvulaceae | \nLC | \n✓ | \n\n | \n |
24 | \nLablab purpureus (L.) Sweet | \nShim | \nFabaceae | \nLC | \n✓ | \n\n | ✓ | \n
25 | \nLagenaria siceraria (Molina) Standl. | \nLau | \nCucurbitaceae | \nLC | \n✓ | \n\n | ✓ | \n
26 | \nLuffa acutangula (L.) Roxb. | \nJhinga | \nCucurbitaceae | \nLC | \n✓ | \n\n | ✓ | \n
27 | \nLuffa cylindrica (L.) M. Roem. | \nPurul | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
28 | \nMikania cordata (Burm. f.) Robinson | \nAssamlata | \nAsteraceae | \nLC | \n\n | \n | ✓ | \n
29 | \nMomordica charantia L. var. charantia C. B. Clarke | \nKarolla | \nCucurbitaceae | \nLC | \n✓ | \n\n | ✓ | \n
30 | \nMomordica cochinchinensis (Lour.) Sprengel | \nKakrol | \nCucurbitaceae | \nLC | \n✓ | \n\n | ✓ | \n
31 | \nPiper betle L. | \nPan | \nPiperaceae | \nLC | \n✓ | \n\n | \n |
32 | \nSmilax perfoliata Lour. | \nKumari lata | \nSmilacaceae | \nLC | \n\n | \n | ✓ | \n
33 | \nSolena amplexicaulis (Lamk.) Gandhi | \n\n | Cucurbitaceae | \nLC | \n✓ | \n\n | \n |
34 | \nTapiria hirsuta Hook. f. | \n\n | Anacardiaceae | \nCD | \n✓ | \n\n | \n |
35 | \nTetrastigma bracteolatum (Wall.) Planch. | \nGolgoli lata | \nVitaceae | \nCD | \n\n | \n | ✓ | \n
36 | \nTrichosanthes anguina L. | \nChichinga | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
37 | \nTrichosanthes dioica Roxb. | \nPatal | \nCucurbitaceae | \nLC | \n✓ | \n\n | \n |
38 | \nUvaria hirsuta Jack | \nBanor kola | \nAnnonaceae | \nNE | \n\n | \n | ✓ | \n
39 | \nVigna unguiculata (L.) Walp. | \nBorboti | \nFabaceae | \nLC | \n✓ | \n\n | \n |
40 | \nAngiopteris evecta (Forst.) Hoffm | \nDhekia Shak | \nAngiopteridaceae | \nLC | \n\n | ✓ | \n\n |
41 | \nBlechnum orientale L. | \n\n | Blechnaceae | \nLC | \n✓ | \n✓ | \n\n |
42 | \nChristella arida (D. Don) Holtt. | \nBish Dhekia | \nThelypteridaceae | \nLC | \n\n | \n | ✓ | \n
43 | \nDiplazium esculentum (Retz.) Sw. | \nDhekia, Dhekia Shak | \nAthyriaceae | \nLC | \n✓ | \n✓ | \n\n |
44 | \nHelminthostachys zeylanica (L.) Hook. | \nShada Dhekia | \nHelminthostachyaceae | \nLC | \n\n | ✓ | \n✓ | \n
45 | \nLygodium microphyllum (Cav.) R. Br. | \n\n | Lygodiaceae | \nLC | \n✓ | \n\n | \n |
46 | \nMarsilea quadrifolia L. | \n\n | Marsileaceae | \nLC | \n✓ | \n\n | \n |
List of fodder yielding climbers and ferns recorded from the protected areas [here, CD, conservation dependent; LC, least concern; NE, not evaluated; NE (rare), not evaluated but seems to be rare; NT, near threatened; V, vulnerable].
Threats to the fodder yielding plant species are similar to that of the protected areas and forests across the different regions of Bangladesh. According to the fifth report on CBD submitted by the Bangladesh government in 2015, direct threats to the PAs are (1) encroachment in protected areas, (2) degradation of forests and wetlands, (3) infrastructure development, (4) unsustainable and/or illegal exploitation of terrestrial resources, (5) unsustainable and/or illegal fishing practices, (6) change in hydrological regime, (7) pollution, and (8) invasive species [33]. On the other hand, the indirect threats are the institutional and environmental conditions that are behind the direct threats visible on the ground [34]. The main indirect threats are (1) poor institutional capacity, (2) lack of coordination among different agencies, (3) policy and information gaps, (4) lack of enforcement, (5) inadequate and poorly managed system of protected areas, (6) corruption, (7) lack of political commitment, (8) lack of awareness, (9) climate and biophysical changes, and (10) lack of alternative livelihoods in sensitive habitats.
\nAn unprecedented threat to the fodder plant diversity of Bangladesh is exerted by the conversion of wildlife habitat into human settlements along with rampant urban development throughout the country’s forested areas [35].
\nThe ecosystem integrity of the PAs of Bangladesh are in very vulnerable situation because they are part of reserved forests which have, in most cases, only been declared after being degraded heavily by various means. It has been assumed that 10% of it is already extinct due to overexploitation. The Bangladesh National Herbarium (BNH) has reported 106 vascular plant species at varying degrees of risk of extinction [33].
\nThe main threats to flora and fauna of protected area conservation emanate from the degradation of forests and wetlands. It is assumed that the anthropogenic pressures on natural resources caused ecosystem depletion. Biotic pressures are exacerbated by dramatic change in climate pattern coupled with sea level rise, increase soil salinity, and increase incidence and severity of cyclones and change in rainfall patterns and temperatures, disturbing the regular seasonality of fruit and flower blooms. This impacts the regeneration of important flora and fauna species and disrupts food chain.
\nA study revealed that protected areas in the tropical moist evergreen and semievergreen forests of hilly regions were highly subject to illegal wood cutting, while those in tropical moist deciduous forests of plain land area were prone to encroachment for settlement and agriculture, and those in mangrove forests of littoral zones were extremely vulnerable to wildlife poaching [36].
\nPlants and animals are two of the main interacting components of an ecosystem. A very close symbiotic relationship exists between flora and fauna including microbes, i.e., fungi, algae, bacteria, etc. Pollination, decomposition of degradable wastage, nutrient cycling, forming food web, and maintaining the food chain are the main fields where contribution of fauna is very significant. On the other hand, supplying nutrients through food production, providing shade and shelter, and maintaining fertility and productivity of an ecosystem are the key contributions of the floral communities to the associated ecosystem. The smaller animals use the cover of plants and dead leaves to hide from the predators. These animals, i.e., moth, katydids, frogs, and grasshoppers, can blend into the surrounding environment at will and become invisible to the predators searching for food. The tropical rain forests like the protected areas of Bangladesh are very much responsive to animal and plant interaction. In adaptive surroundings of tropical forests, a huge diversity of animals, birds, and insects subsist together. An area of 6 square kilometer of typical tropical rain forest can harbor as much as 1500 flowering plants, 400 species of birds, 100 reptiles, and 60 amphibians along with thousands of butterfly species. However, in a complex ecosystem, the biotic interactions can be toward any directions, i.e., plant-plant, plant-animal, plant-microbe, animal-microbe, and animal-animal. All of these interactions employ different biotic services [37].
\nThe richness of a wildlife habitat with fodder very sharply determines the fluctuation of wildlife population. Unavailability of food inside the forests make many of the wildlife to come out toward adjacent localities in search of food. For example, a higher number of human-elephant conflicts (HEC) were reported from the southeastern Bangladesh due to degradation and fragmentation of elephant habitat which caused reduction of Asian elephant’s population from 500 in the middle of the last century to 228–327 [38]. It is worth and interesting to mention that in some protected areas like Nijhum Dweep National Park, the deer population is shrinking gradually due to higher competition with the thousands of buffalos and cows for fodder. Similar to that of the Asian elephant and deer, the population of monkey along with other herbivores is also reducing at alarming rate due to degradation of overall habitat quality in Bangladesh [39, 40].
\nThe process of conserving rare plant species can be divided into three phases: (i) Identification-determining which species are in danger of extinction. (ii) Protection-determining and implementing the short-term measures necessary to halt a species’ slide to extinction. (iii) Recovery-determining and implementing the longer-term measures necessary to rebuild the population of the species to the point at which it is no longer in danger of extinction [41].
\nPeople living in and around a forest depend on forest resources for a substantial proportion of their subsistence, including food, fiber, medicines, and other uses [42]. Many others perceive forest exploitation as a means of escaping poverty [43, 44]. Forest conservation is likely to be low on these peoples’ priorities if it limits their possibilities for livelihood support [45]. So, the development of living condition of the surrounding local people of the important wildlife habitats through improving their livelihood security and diversifying income, so as to meet all their basic needs, should be of first priority; otherwise the conservation effort will go in vain. It is important to extend and strengthen the protective measures by local administrative bodies of Bangladesh Forest Department (FD) against the threats like settlement, agricultural expansion, severe grazing, hunting, shooting, illegal cutting, etc. Local administrative units (beat offices) of the Forest Department must be strengthened with necessary manpower, staff quarters, equipment, logistics, and training, so that they become more capable to conduct the protection and conservation measures because they are the only authority to look after forest and wildlife.
\nSteps should be taken to halt further expansion of any agricultural/horticultural practices toward the forests. Awareness raising and consciousness of local people regarding the importance of habitat integrity, wildlife, environmental conservation, biodiversity, and endangered ecosystems are mandatory. The government may provide some incentives through money, small loan, training, etc. to help local people in managing sustainable alternative income-generating programs. Relocation and permanent allocation of some barren lands to the forest-dwelling people may reduce their dependency and threats induced by them on the existing forest. Cultivation of crops, i.e., pineapple, banana, paddy, taro, and lemon, should be restricted to some marginal areas of the forests or protected areas. Cattle grazing and browsing pressure in many protected areas is significant, and hence control of grazing animals for reducing the direct effects of disturbance is essential. Special conservation measures both ex situ and in situ methods may be initiated to conserve the threatened and rare native plant species. Enrichment plantation with native wildlife (i.e. rhesus macaque, capped langur, etc.) fruit-bearing plant species should be conducted in the gap spaces of the forests. Activities that were identified by the IUCN and different conservation organizations which contribute to forest and species conservation, i.e., area-based protection, area-based management, species-centered management, education and awareness, improved law and policy, livelihoods and incentives, and capacity building, may be considered for conservation of forest resources in all protected areas. Permanent sample plots of adequate size (0.5–1.0 ha) may be established in representative vegetation types of each wildlife habitat to facilitate long-term ecological and biodiversity assessment which may help monitor the success of restoration and conservation of the fodder yielding plants. Community patrolling should be strengthened to reduce illicit felling as well as raise awareness among local people regarding nature conservation. Digging furrow in suitable locations across the forest may be helpful for controlling fire infestation and litter extraction by local people using small vehicles.
\nFragmentation and degradation of wildlife habitat quality resulted in reduction of the diversity and population of fodder yielding plants. It is a worldwide trend in which the policy makers and scientists are concerned about. However, Bangladesh having a substantial area declared as protected areas is trying to conserve and restore the wildlife habitat quality. Still now, the protected areas of the country are still harboring a good number of fodder yielding plants from all habit forms. However, many of the fodder species are having very poor density which may reduce further and get extinct if appropriate species-specific multiplication and conservation measures are not taken immediately.
\nThe authors acknowledge the contribution and supports of the Bangladesh Forest Department, Arannayk Foundation, and University of Chittagong during the studies.
\nIn this chapter we will analyze the physiopathological changes involved in the inflammatory response of the septic process in infective endocarditis [IE] that culminate with cellular damage and the generation of organic failures; morphological changes, cellular biology, biochemistry, immunology, and genetic vulnerability, which together are called “pathobiology,” are the substrate of clinical manifestations of this serious disease, which requires a multidisciplinary group of experts (cardiologists, infectologists, surgeons, intensivists) to optimize therapeutic approach. IE is defined as a severe multisystem disease, which results from an infection, often bacterial, that initially affects the endocardial surface of the heart [1]. The epidemiological pattern has changed over time [2, 3, 4]. The incidence has increased in recent years to 5–10 cases per 100,000 inhabitants [2], due to the fact of a greater number of predisposing factors such as the use of permanent cardiovascular devices, invasion with intravenous catheters in critical care units, and hemodialysis treatments, in addition to having greater accessibility to diagnostic tools. From the etiological point of view, Staphylococcus aureus (S. aureus) is predominant as a causal germ [5, 6]. The clinical course of a patient with IE depends on the inflammatory response, since it is variable; it also depends on the germ and the response of the patient to infection with varying degrees of hemodynamic and metabolic compromise [7, 8, 9]. We emphasize the current trend of the search for organic failures associated to the septic processes for their identification and stratification and therapeutic approach [10]. Given the characteristics of the disease, IE has a high mortality that goes from 20 to 30% in the reported series [2, 11]; it is noteworthy that the evolution toward septic shock has been documented in 30% [12], considering this complication as an independent variable of poor prognosis [13].
The pathogenesis and the prognosis of IE can be simply described in a general way as the interaction between the host and the germ; however, these factors are not independent and are very importantly linked both in the susceptibility characteristics of the host (advanced age, higher prevalence of comorbid conditions, and exposure to health care) to survive or not to an infectious state, as of the characteristics of the germ involved. To reduce the incidence of IE and improve its outcome, epidemiological studies can provide valuable information on contemporary and modifiable risks to modify their morbidity and mortality [14].
The incidence of hospital discharge diagnoses for drug dependence combined with IE increased more than twelvefold from 0.2 to 2.7 per 100,000 persons per year over this 6-year period. Correspondingly, hospital costs for these patients increased eighteenfold, from $1.1 million in 2010 to $22.2 million in 2015 [15].
In another study also conducted in the USA, using a national sample of hospitalized patients from 1998 to 2009 with focus on IE showed an increase in the use of intracardiac devices from 13.3 to 18.9%. In cases with pathogens identified, S. aureus was the most common, increasing from 37.6% in 1998 to 49.3% in 2009, 53.3% of which were methicillin-resistant Staphylococcus aureus (MRSA) [16]. The above can give us an idea of the economic and assistance impact of treating patients with severe sepsis such as IE. It is an infection inside the organ that is responsible for distributing blood to practically the whole organism.
The evolution of an inflammatory process plus infection frequently occurs with clinical manifestations unspecified such as fever or hypothermia, tachycardia, tachypnea, or abnormal white blood cell count, progressing to septic shock and acute organ failure [17].
Epidemiological data of more than five decades tell us that S. aureus is the most important causal agent of IE [4]; so in the development of systemic inflammation that is generated by the host-germ interaction, we will consider the S. aureus as the best example of IE due to its virulence and an emergent property that we know as resistance to antibiotics, sophisticated defense mechanisms, and the ability to cause apoptosis in cells when it is alive inside the cell. The interaction of S. aureus-host allows us to develop in a substantive way, on one hand, the importance of the virulence of the germ and, on the other, the defense mechanisms of the host, showing how the inflammation is generated and amplified to offer a step to oxidative stress. It is important to mention that other agents can cause IE such as streptococci and fungi.
S. aureus is a Gram-positive coccus with a diameter of 0.5–1.5 μm, grouped as single cells, in pairs, tetrads, short chains, or forming a conglomerate in a cluster of grapes. This microorganism was first described in the year 1880, in Aberdeen, Scotland, by the surgeon Alexander Ágoston. The name comes from the Greek σταφυλόκοκκος, which is composed of the terms “staphylé,” meaning cluster, and coccus, meaning grain or grape, and from the Latin “aureus” which means golden, that is to say “cluster of golden grapes.”
They are non-motile bacteria, not sporulated, with no capsule (although there are some strains that develop a slime capsule); they are facultative anaerobes. Most staphylococci produce catalase (enzyme capable of dismutating hydrogen peroxide in H2O + O2), characteristic that is used to differentiate its sort from others like Streptococcus and Enterococcus. In 1961, the first report was made on the existence of a methicillin-resistant Staphylococcus aureus [18].
S. aureus is a pathogen that causes significant morbidity and mortality worldwide [2]. It is the leading pathogen associated with life-threatening bloodstream infections [19].
Although S. aureus is mainly known as an extracellular pathogen, it has been shown to invade and survive within endothelial cells, both within vacuoles and free in the cytoplasm, which implies that the bacteria can escape from the phagolysosome. S. aureus tends to infect endovascular tissue. It is believed that this ability contributes to causing a persistent endovascular infection with endothelial destruction.
On the other hand, the death of endothelial cells after the invasion of S. aureus occurs at least in part by apoptosis, as demonstrated by DNA fragmentation and changes in nuclear morphology. Apoptotic changes are observed as early as 1 h after infection of endothelial cells [18]; they are considered to function as nonprofessional phagocytes, being able to ingest S. aureus [20, 21] following the adhesion of this to endothelial cell monolayers; invasion can occur through ingestion by endothelial cells.
For the internalization of S. aureus, adherence seems to be necessary, since the use of the phagocytosis inhibitor cytochalasin D prevented apoptosis. Studies show that living intracellular S. aureus induces apoptosis of endothelial cells and that this depends on a factor associated with viable organisms, since dead S. aureus (by ultraviolet light) also internalized does not induce it [18]. The process has been observed through electron transmission micrographs of bovine aortic endothelial cell monolayers infected with S. aureus, showing phagocytosis following a sequence of events: (I) adhesion of S. aureus to the endothelial cell, (II) formation of cup-shaped processes on the surface of the endothelial cell underlying the adherent bacteria, and (III) elongation of the cup and engulfment of bacteria within a phagosome [19].
To colonize a vertebrate host, S. aureus requires numerous nutrients, such as the prosthetic group heme. The requirement can be met through two distinct mechanisms: importing exogenous heme through dedicated machinery or synthesizing endogenous heme from own metabolic precursors. These two mechanisms are necessary for a complete virulence of S. aureus [22, 23]. Once acquired, heme is used for several cell processes. The intact heme is used as a cofactor for enzymes [24], including cytochromes in the electron transport chain, catalase for the detoxification of reactive oxygen species, and bacterial nitric oxide synthase (bNOS).
Although the S. aureus requires heme, its excess is toxic to the germ, so it has a mechanism for hem detoxification through a hem sensor system (HssRS) that induces the expression of a hem regulator transporter (hrtAB) [25]. The suppression of the components of this route affects the virulence of S. aureus. This ability to detoxify heme is critical to survive in the host. Also, the synthesis of nitric oxide is important for the bacteria to survive. Bacteria encode genes similar to nitric oxide synthetase in mammals, which leads to the characterization of the nitric oxide synthase hemoprotein (bNOS) [26].
The S. aureus contains molecules such as peptidoglycan and lipoteichoic acid, potent stimulants for the production of cytosines such as TNF-α, IL-4, IL-6, IL-8, IL-12, IL-1be, growth-regulated oncogene (GRO) alpha, and regulated upon activation, normal T-cell expressed, and secreted (RANTES). RANTES has a chemotactic function to perform leukocyte recruitment to areas of infection in addition to inducing tumor necrosis factor alpha and interleukin 1. Elevated levels can persist for 7–14 days [27]. As we can observe, S. aureus activates in a very important way the process of inflammation.
Circulatory blood stream infections (positive blood cultures) occur in patients with intravascular prosthetic devices as the most common source of infections related to health care [28]. MRSA was the most frequent pathogen in these types of infections with a consistent increase in the isolates of MRSA [29, 30, 31]. In the EU, epidemiological surveillance data on bloodstream infections show a marked variability among the member countries that make up a proportion of S. aureus that is resistant to methicillin, ranging from less than 1% to more than 50%. In addition to infections associated with health care, new MRSA strains have emerged in their communities as human pathogens associated with livestock [32].
The anatomical and physiological barriers of cardiac protection such as the endothelium can be compromised in its structure when areas of turbulence and injury are generated, producing an area exposed to infection. The intracardiac cavities have a cell layer called endocardial endothelium (EE) that covers the endocardium of the atria, ventricles, and all their anatomical components (papillary muscles, chordae tendineae, and heart valves). The EE acts as an active mechanism of biological heart-blood barrier, since it interacts dynamically with cardiomyocytes allowing direct communication and signaling between both types of cells. This electrochemical communication between the cells of the EE and the cardiomyocytes allows a rapid intracellular electrochemical propagation and amplification of the functional properties of the EE.
Signaling between cardiac endothelial cells (EE and myocardial capillary endothelium) and cardiomyocytes influences cardiac growth, contractile performance, and rhythmicity. The network of Purkinje fibers and the subendocardial neural plexus (parasympathetic nervous system) is immediately below the endocardial endothelium (EE) and participates in the endothelial control of cardiac rhythm. Endothelin-1 (Et-1), nitric oxide (ON), prostaglandins (PGI2), prostacyclin (AI and AII), angiotensin I and II, and vascular endothelial growth factor (VEGF) are involved in these processes.
The endothelium that covers cardiac structures is at the vascular level, the myocardial capillary, and the endocardium; its activation includes changes in the endothelial phenotype as part of the physiological adaptive response to several possible injuries and stressors. The dysfunction of the endothelium implies a deregulated response that is not useful and that can be permanent.
One of the clinical disorders that selectively damage the endocardium and subendocardial interstitial tissue is endocarditis. This entity causes activation of the vascular and endocardial endothelial system, as well as poor adaptation or failure characterized by hemodynamic abnormalities, neurohormonal imbalance, cytokine expression, and endothelial dysfunction [33].
Infective endocarditis is an anatomoclinical entity characterized by microbial infection of the valvular or parietal endothelium or both; it is located predominantly on the left side of the heart, although it can also occur in the right (e.g., endovenous drug), which produces inflammation, exudation, and proliferation of the endocardium. The most characteristic lesion is the vegetation, constituted by an amorphous mass of platelets and fibrin, of variable size, which contains multiple microorganisms and scarce inflammatory cells (fibrinoplaquetary thrombus) [34]. This type of lesions generates metastatic infection in other anatomical territories, for example, the central nervous system, apostematous meningitis, myocarditis, pyelonephritis, and splenic abscesses which are at risk of rupture [35, 36].
The clinical manifestations of infective endocarditis are acute rapidly progressive or subacute; the pathophysiological processes of both are explained by immunological and vascular phenomena, such as inflammatory response, mediators of inflammation triggered by a maladaptive response to an infectious process, aggregation of immune complexes, infectious vasculitis, and peripheral microembolism [34, 37]. Depending on the affected cardiac cavity (right/left) or valvular system, the clinical manifestations will be due to the aforementioned processes [38] (Table 1).
Patients, % | |
---|---|
Sign | |
Fever | 86–96 |
New murmur | 48 |
Worsening of old murmur | 20 |
Hematuria | 26 |
Vascular embolic event | 17 |
Splenomegaly | 11 |
Splinter hemorrhages | 8 |
Osler nodes | 3 |
Janeway lesions | 5 |
Roth spots | 2 |
Complication | |
Stroke | 17–20 |
Non-stroke embolization | 23–33 |
Heart failure | 14–33 |
Intracardiac abscess | 14–20 |
New conduction abnormality | 8 |
The anatomopathological changes due to the formation of vegetations in the valvular ring and/or in the leaflets cause an anatomical alteration. If this anatomical alteration generated by a vegetation prevents valvular closure, it will be expressed as a murmur of valvular insufficiency and in severe cases such as microembolisms septic and non-septic and cardiac failure [34, 37].
The standard reference to corroborate the clinical diagnosis of IE is transesophageal echocardiography since the transthoracic echocardiogram, even when limited to native valves, decreases the diagnostic probability of IE [39].
Right and left endocarditis are two distinct entities that require different clinical and surgical approaches. The diagnosis of endocarditis on the right side requires a high index of clinical suspicion. It can occur with a history of intravenous drug use, fever, and pulmonary infiltrates, although intravenous drug abuse is also a cause of IE on the left side of the heart [36]. The information provided by echocardiography is of prognostic and therapeutic value.
If the vegetation is <1.0 cm in diameter, it can be expected that antibiotic therapy will resolve the infection; if the size of the vegetation determined by echocardiography is ≥1.0 cm without response to treatment, surgical intervention should be considered [40].
Surgical treatment in IE on the left side of the heart, for example, the mitral valve, is indicated in patients with severe mitral regurgitation, even in the absence of congestive heart failure, with mitral annular abscess, large vegetation >10 mm, uncontrolled sepsis, and multiple embolisms [41]. Mitral valve (MV) replacement has traditionally been considered as the standard treatment for MV endocarditis that does not respond to antibiotic treatment.
However, the pioneering work of Dreyfus et al. surgery for repair of the mitral valve with IE can be performed safely and is often associated with a better outcome compared to mitral valve replacement [42, 43].
It has been largely recognized that infective processes have considerably different patient-to-patient behavior in such a way that some patients respond well to the treatment applied and some others end up developing a dysregulated immune response known as sepsis [44], organ failure, and some even die from this process. Infective endocarditis does not escape from this fact. Many variables, such as the virulence of the pathogen and the quality of the treatment applied, among many others, participate in an additive manner to conform the clinical outcomes of infections, and this helps to understand why a patient takes the road of success or failure regarding the control of the septic process. One of the most recent advances in the understanding of the pathophysiology of infective processes, including infective endocarditis, is the demonstration that genetically determined differences in the immune system of individuals are one of these many factors that determine the phenotypic behavior and outcomes. Therefore, the next chapter section is dedicated to explaining the existing evidence of the participation of immunogenetics in the development of infective endocarditis and sepsis.
Recently, the concept of sepsis has been redefined as the result of a better understanding of its pathophysiology, particularly regarding the early activation of pro- and anti-inflammatory immune responses. As the third international consensus definition of sepsis states, sepsis is a life-threatening organ dysfunction caused by a dysregulated host response to an infection [45]. Then, if sepsis is dependent on a dysregulated response, and this response is executed principally by the host immune system, then genetically defined differences between individuals immune system might play a role in the genesis of this syndrome and at least partially explain why some patients take the road of sepsis and some others do not. This hypothesis had long been existed, but it was until 1988 that the theory started gaining scientific evidence of its existence, when Sørensen et al. [46] published what is considered a landmark study with respect to this topic. In this article, the authors studied the genetical influences on the principal causes of nonviolent premature death in the Danish population; to separate them from the environmental influences, they studied a selected group of people that had been adopted early in life. This was extracted from the Danish Adoption Registry and included adoptees that were born between 1924 and 1926. They traced them up and demonstrated that the death of a biologic parent from an infection before the age of 50 resulted in a relative risk of death from infective causes in the adoptees of 4.5. Since this publication, a great number of studies have been conducted in an attempt to define the specific genetic variations that determine these differences in outcome. This task has resulted complex; as both pro- and anti-inflammatory responses contribute to the outcome of septic processes, all genes encoding effector proteins in the biochemical pathways of the inflammatory response to infection are potential candidates to determine the genetical background responsible for the interindividual differences aforementioned [47].
5.1.1 The study of single nucleotide polymorphism associations with sepsis and IE outcomes.
The most studied specific type of genetic anomaly regarding to sepsis susceptibility is the single nucleotide polymorphism (SNP); therefore, the largest body of evidence comes from the study of this type of genetic variations. SNPs are defined as frequent (occurring in >1% of the population) variations in the human DNA sequence [48] and consist in the exchange of a single base pair for another in a specific location in the DNA sequence. They may occur within the exonic (coding) or intronic (noncoding) region of the gene and can have different consequences which include alteration of expression or structure of proteins and enzymes, introduction of an alternative translation initiation codon or stop codon, and destabilization of exonic mRNA [49]. Methodologically speaking, most studies are association studies (case/control and cohort type), and two approaches have been done. In the most common approach (which for purposes of this chapter section are going to be called specific SNP association studies), the frequency of one or more known SNPs present in genes coding defined molecular candidates involved in the pro- or anti-inflammatory responses (e.g., alpha tumoral necrosis factor gene) is compared between a specific phenotypically defined interest group (patients with a confirmed specific infectious scenario as sepsis or IE) and a control group, usually consisting of a group of healthy blood donors ideally with an ethnicity equal to the interest group. If there are statistically significant differences in the frequency of the SNPs between groups, authors take this as proof that such genetic differences are implicated in the specific way that the study population responds to infection. The other approach is a type of study called genome-wide association studies (GWAS). As the previously described type of study, GWAS are association studies (most frequently case–control studies) but differ in that the frequency of most known SNPs is measured in the whole genome of the cases (infected group) and controls (healthy blood donors). When a statistically significant difference is found, authors take this as proof that such genetic variability is responsible for the difference in outcomes and then hypothesize, based in the location of the SNPs, about the biological plausibility of the association given the gene that is affected.
A large number of specific SNP association studies have been conducted respecting the most important effector molecules in response to sepsis and also some GWAS.
In response to an infectious stimuli, such as lipopolysaccharides (LPS), tumor necrosis factor α (TNF-α) is a cytokine that is released early mainly by macrophages, and it is a principal mediator of the inflammatory response to infection which stimulates acute inflammation by its action on different cells, such as endothelial cells and leukocytes [50].
Many studies have been done in an attempt to determine if specific SNPs in the TNF alpha factor gene are implicated in sepsis susceptibility with conflicting results. A recent meta-analysis from Zhang et al. [51] which included 23 articles that evaluated the effects of TNF-α rs1800629 and rs361525 polymorphisms on sepsis risk found that TNF-α rs1800629 was associated with increased sepsis risk in the overall population in four genetic models, including adenosine (A) vs. guanine (G) (p < 0.001, odds ratio (OR) = 1.32), GA vs. GG (p < 0.001, OR = 1.46), GA + AA vs. GG (p < 0.001, OR = 1.46), and carrier A vs. carrier G (p < 0.001, OR = 1.32). These results suggest an implication of these genetic variations with an increased susceptibility for sepsis development.
TNF-β is a cytokine produced by T lymphocytes similar to TNF-α and binds to TNF receptors. It activates endothelial cells and neutrophils and is a mediator of acute inflammatory response, providing a link between T-cell activation and inflammation. These effects are the same as those of TNF-α, consistent with their binding to the same receptors. However, as the quantity of TNF-β is much less than that of TNF-α made by lipopolysaccharide-stimulated mononuclear phagocytes, TNF-β is not readily detected in the circulation. For this reason, TNF-β is usually a local cytokine and not a mediator of systemic injury. A single nucleotide polymorphism has been found at position +252 in the first intron of the TNF-β gene and consists of a G in the wild-type allele (TNFB1) and an A in the variant allele (TNFB2). Known as the Nco1 polymorphism, it has been proposed as a potentially influential locus in many inflammatory conditions. Delongui et al. studied the association of the TNF-β Nco1 genetic polymorphism with susceptibility to sepsis in 60 patients diagnosed with sepsis and in 148 healthy blood donors. Among the septic patients, the allelic frequencies of TNFB1 and TNFB2 were 0.2833 and 0.7166, respectively, and they differed from those observed in the blood donors (p = 0.02). The TNFB2 allele frequency was higher in the septic patients than in the controls [OR = 1.65 (CI 95% 1.02–2.69), p = 0.0315], all this suggesting an implication in susceptibility to sepsis [52].
IL-10 has beneficial anti-inflammatory properties; however, an excess of IL-10 has been reported to induce immunosuppression in bacterial sepsis. Published data demonstrates that lower production of IL-10 from stimulated peripheral blood mononuclear cells (PBMC) from septic patients is significantly correlated with favorable disease outcome [53]. Stanilova et al. [54] investigated the −1082 (A/G) polymorphism in the promoter of the IL-10 gene by measuring IL-10 production from stimulated peripheral blood mononuclear cells (PBMC) and to evaluate the relationship of this polymorphism with susceptibility to severe sepsis and its outcome. They found that carriage of at least one copy of IL-10-1082 G allele in sepsis patients and in healthy controls resulted in a statistically significant increase in IL-10 production from stimulated PBMC. Patients who survived sepsis had a significant decrease of IL-10-1082 allele G frequency, compared with controls (17 vs. 47.2%; p = 0.012). This suggests that this genetic variation has an impact in IL-10 production and in the outcomes of septic patients [55].
Interleukin 1β (IL-1β) is a potent pro-inflammatory cytokine implicated in the development of chronic inflammatory disorders. IL-1β signaling is blocked by IL-1 Ra, a natural regulator of IL-1 cytokines. IL-1 Ra binds to the IL-1 receptor and thereby prevents binding of both IL-1a and IL-1b [56]. F. Arnalich et al. aimed to determine the influence of the polymorphism within the intron 2 of the IL-1RNa (IL-RNa*) on the outcome of severe sepsis. A group of 78 patients with severe sepsis (51 survivors and 27 non-survivors) was compared with a healthy control group of 130 blood donors and 56 patients with uncomplicated pneumonia. They found a significant association between IL-1RN* polymorphism and survival. After adjusting for age and APACHE II score, they did a multiple logistic regression analysis that showed that patients’ homozygotes for the allele *2 had 6·47 times more risk of death (95% CI 1·01–41·47, p = 0.04). These authors concluded that these genetic mutations might be implicated in an increased risk of death in septic patients [57].
HMGB1 is a pleiotropic cytokine that has been implicated in the pathophysiology of systemic inflammatory response syndrome (SIRS) and sepsis. HMGB1 is measurable in the systemic circulation in response to severe injury. This protein has the propensity to bind to a variety of inflammatory mediators such as lipopolysaccharide and pro-inflammatory cytokines, including IL-1. The role of HMGB1 as an endogenous molecule facilitates immune responses and has an important role in homeostasis between tissue and disease. HMGB1 is implicated in the pathophysiology of a variety of inflammatory diseases, and it has been found that variation in the HMGB1 gene is associated with mortality in patients with systemic inflammatory response syndrome [58]. Kornblit et al. performed a long-term, 4-year study comparing HMGB1 sequencing data in 239 intensive care unit (ICU) patients with HMGB1 blood levels and clinical outcomes. The promoter variant −1377delA was associated with a markedly reduced long-term survival rate after ICU admission in SIRS patients. There was also a significant interaction with a polymorphism within the coding region of the HMGB1 gene at position 982 (C > T) in exon 4; carriers had an increased frequency of early death from infection [59].
TLRs are a group of pattern recognition receptors. They play important roles in regulating inflammatory reactions and activating adaptive immune response to eliminate infective pathogens [60]. TLR2, a key member of TLR family, can recognize a variety of bacterial lipoproteins. The mechanism of TLR2-recognizing lipoproteins has been elucidated; after TLR2 recognizes lipoproteins, it activates MyD88 adaptor-like protein and initiates a signaling pathway, which induces further immune response [61]. This evidence puts TLR2 gene as an appealing candidate for determining sepsis risk. In a recent meta-analysis, Gao et al. [62] analyzed a total of 12 studies (11 records) with 898 cases and 1517 controls examined to determine the association between the TLR2 Arg753Gln polymorphism and sepsis risk. The combined results of the overall comparison indicated that there were significant associations between the TLR2 Arg753Gln polymorphism and sepsis risk under the allele comparison model and the dominant model, respectively (for A vs. G, OR 1.76, 95% CI 1.05–2.95, p = 0.03; for AA/GA vs. GG, OR 1.92, 95% CI 1.11–3.32, p = 0.02).
Rautanen et al. [63] did a genome-wide association study in three independent cohorts of white adult patients admitted to ICU with sepsis, severe sepsis, or septic shock due to pneumonia or intra-abdominal infection (n = 2534 patients). The primary outcome was 28-day survival. Results for the three cohorts of patients with sepsis due to pneumonia were combined in a meta-analysis of 1553 patients. The most significantly associated SNPs were genotyped in a further 538 white patients with sepsis due to pneumonia (an independent fourth cohort), of whom 106 died. In the genome-wide meta-analysis of three independent pneumonia cohorts, common variants in the FER gene were strongly associated with survival (p = 9·7 × 10–8; OR 0·52 [95% CI 0·41–0·66]). Genotyping of the additional fourth cohort strengthened the evidence for association with survival (p = 5·6 × 10–8; OR 0·56 [0·45–0·69]).
There are many risk factors described for the development of IE; nevertheless up to 30–50% of patients with this diagnosis does not have any known risk factor [64]. Therefore, as in sepsis per se, there is thought to be immunogenetic influences that affect the risk of development and outcomes in IE. However, in comparison to sepsis, evidence of the immunogenetic influence on the susceptibility and outcomes of IE is less robust. Golovkin et al. [65] hypothesized that inherited variation in TLR and triggering receptor expressed on myeloid cells (TREMs) genes may affect individual susceptibility to IE. They conducted a specific SNP study in which the distribution of genotypes and alleles of the TLR1, TLR2, TLR4, TLR6, and TREM-1 gene polymorphisms was investigated in 110 Caucasian subjects with IE and 300 matched healthy blood donors. ORs with 95% CI were calculated. They found that C/C genotype of the rs3775073 polymorphism within TLR6 gene was associated with a decreased risk of IE (OR = 0.51, 95% CI = 0.26–0.97, p = 0.032) according to the recessive model; however, there was no association between the other investigated SNPs within TLR andTREM-1 genes and IE.
Moreau et al. [66] conducted a GWAS of 67 patients with definite native valve S. aureus IE (cases) and 72 matched native valve patients with S. aureus bacteremia but without IE (controls). Unfortunately, no SNPs were significantly associated with S. aureus IE at the genome-wide level (p < 5 × 10−8). Four suggestive SNPs (p < 0.00001) were located on one locus on chromosome 3, near the genes CLDN11 and SLC7A14. For all, the frequency of the minor allele was lower in cases than in controls, suggesting a protective effect against S. aureus IE. The same association was observed using an independent Danish verification cohort of Staphylococcus aureus bacteremia with (n = 57) and without (n = 123) IE. An ex vivo analysis of aortic valve tissues revealed that S. aureus IE associated SNPs mentioned above were associated with significantly higher mRNA expression levels of SLC7A14, which is a cationic amino acid transporter protein. These results suggest an IE-protective effect of SNPs on chromosome 3 during S. aureus bacteremia. The authors concluded that the effects of protective minor alleles may be mediated by increasing expression levels of SLC7A14 in valve tissues.
The modern mitochondria have an evolution of more than a billion years, originating as an invading Eubacterium in early eukaryotic cells. The knowledge of the structure, functionality, and the similarities of the DNA between mitochondria and bacteria strongly prove the endosymbiotic origin of the mitochondria. Of the 1000 or more mitochondrial proteins, only 13 are encoded by the mitochondrial genome, the rest is transcribed and translated into the nuclear genome and transported to the inner mitochondrial membrane [67].
In the heart the populations of mitochondria include subsarcolemmal mitochondria, which are more susceptible to injury. Subsarcolemmal mitochondria provide energy for membrane-related processes, including signal transduction, ion exchange, and substrate transport, whereas the intermyofibrillar mitochondria more directly support muscle contraction [68].
The mitochondrial oxidative phosphorylation process is responsible for the conversion of macronutrient energy (e.g., glucose, fatty acids, and amino acids) into ATP through a set of coordinated and highly coupled reactions where the macronutrients are oxidized and the oxygen is reduced to water and adenosine diphosphate (ADP) is phosphorylated to ATP.
In the chemiosmotic hypothesis [69], the proton gradient is formed by removing H+ from the interior (matrix), while the negative charges remain inside, largely as OHˉ ions; the pH on the outer face of the membrane (intermembrane space) can reach a pH of 5.5, while the pH just at the inner side (matrix) of the same can reach 8.5; this gradient is 3 pH units. Recall that the pH is equal to - log. of [H+], and therefore 3 units of pH mean that the ΔH+ = 1000 between both faces of the membrane, that is to say there are 1000 times more H+ in the intermembrane space than on the side of the membrane that is in contact with the mitochondrial matrix (Figure 1).
The creation of a proton gradient (∆H+) in the intermembrane space is produced by the chain of electron transport and the synthesis of ATP synthase, which is maintained by the electrons that pass from the reducing equivalents (NADH, FADH2) to the cytochromes along the inner membrane of the mitochondria. ATP synthase uses that gradient to generate ATP. The two processes are associated with the inner membrane of the mitochondria in the mitochondrial crests. Note that the enzymes of the citric acid cycle and β-oxidation are contained in mitochondria, together with the respiratory chain, which collects and transports reducing equivalents, directing them to their final reaction with oxygen to form water, and the machinery for oxidative phosphorylation, the process by which the liberated free energy is trapped as high-energy phosphate. Source: Botham and Mayes [70].
The process begins when carbon substrates enter the tricarboxylic acid cycle through acetyl CoA or anaplerotic reactions. Oxidation of these substrates generates reducing equivalents in the form of NADH and FADH2, which provide electron fluxes through the complexes of the respiratory chain, complex I (NADH dehydrogenase) and complex II (succinate dehydrogenase). The flow of electrons through complexes I and II converges in complex III (ubiquinone-cytochrome c reductase), together with electrons from electron transfer flavoproteins (beta oxidation), although the mobile electron carrier coenzyme Q as second mobile electron carrier transfers electrons to the IV complex (cytochrome c oxidase) where they are finally transferred to oxygen, producing water. A gradient of protons (an electrochemical gradient) through the inner mitochondrial membrane is generated by the action of electron transport through complexes I, III, and IV. The potential energy of this gradient is exploited by the V (ATP synthase) complex to phosphorylate ADP to ATP [71]. It is clear that the maintenance of the mitochondrial membrane potential through the transport of electrons is critical for the proper function of the organelle and, therefore, of the cell and of ascending form of organs and systems.
In the process of mitochondrial respiration, the generation of reactive oxygen species (ROS) is generally a cascade of reactions that begins with the production of superoxide O•2. The oxidative stress is defined as an imbalance that favors ROS production on antioxidant defenses; most ROS are products of mitochondrial respiration. Approximately 1–2% of the molecular oxygen consumed during the process of mitochondria respiration is converted to superoxide radicals. Briefly, the reduction of an electron of molecular oxygen produces a relatively stable intermediate, the superoxide anion (O•2); the importance of this is that it serves as the precursor to most ROS.
Therefore, it is very important to take into account the sources that generate it. There is evidence that most of the O•2 generated by intact mammalian mitochondria in vitro is produced by complex I. The production of superoxide—O•2—is mainly carried out in the inner mitochondrial membrane (IMM) together with complex III [72, 73]. On the other hand, the production of O•2 is stimulated by the presence of succinate (substrate of complex II) [74]. Ubiquinone as part of the respiratory chain binds complexes I with II and II with III which is also important for the formation of O•2 by complex III [75]. Oxidation of ubiquinone—Q cycle—and unstable semiquinone also generates O•2 (Figure 2).
The flavin adenine dinucleotide (FAD) can be reduced in reactions involving the transfer of two electrons (to form FMNH2 or FADH2), but they can also accept one electron to form the semi Quinone. Electron-transferring flavoprotein (ETF). Fe-S, iron?sulfur proteins (nonheme iron proteins). The Fe-S take part in single-electron transfer reactions in which one Fe atom undergoes oxidoreduction between Fe2+ and Fe3+. Coenzyme Q (Q) (also called ubiquinone) (complex I). Cytochrome c, Q-cytochrome c oxidoreductase (complex III), which passes the electrons on to cytochrome c; and cytochrome c oxidase (complex IV), which completes the chain, passing the electrons to O2 and causing it to be reduced to H2O. Q and cytochrome c are mobile. Q diffuses rapidly within the membrane, while cytochrome c is a soluble protein. Mn-SOD, manganese superoxide dismutase.
The Q cycle couples electron transfer to proton transport in complex III electrons are passed from QH2 to cytochrome c via complex III (Q-cytochrome c oxidoreductase) as described in Figure 2.
Superoxide rapidly dismutates into hydrogen peroxide spontaneously or at a low pH is catalyzed by superoxide dismutase. Other elements in the cascade of ROS generation are small molecules derived from oxygen, like the following: hydroxyl (OH•), peroxyl (RO•2), and alkoxyl (RO•) and certain non-radicals that are oxidizing agents and/or are easily converted to radicals, such as hypochlorous acid (HOCl), ozone (O3), singlet oxygen (½O2), and hydrogen peroxide (H2O2). Nitrogen-containing oxidants, such as nitric oxide (NO), are called reactive nitrogen species (RNS), and the Fenton reaction catalyzed by iron leads to the generation of hydroxyl radical [76, 77]. The dismutation of superoxide anions by superoxide dismutases results in the production of H2O2. The mitochondria contribute 20–30% of the stable cytosolic concentration of H2O2 [78]; the subsequent interaction of H2O2 and O•2 in a Haber-Weiss reaction, or the cleavage of H2O2 driven by Fe2+- (or Cu2+), can generate the highly reactive hydroxyl radical (OH•).
The Haber-Weiss reaction [79] may occur as a consequence of oxidative stress. The reaction is catalyzed by the iron in oxidation state (III); the first step of the catalytic cycle is produced by the reduction of the ferric cation to ferrous cation:
The second step is a reaction from Fenton:
Briefly, superoxide dismutases (SOD) are a group of metalloenzymes (containing Fe, Mn, or Cu and Zn) that catalyze the disproportionation of superoxide free radical (2O•) to form hydrogen peroxide and oxygen as shown below:
In some cell types, CuZnSOD is present in the mitochondrial intermembrane space, where it can convert O•2 to H2O2, thus permitting further diffusion into the cytosol.
Superoxide rapidly dismutates into hydrogen peroxide spontaneously or at a low pH is catalyzed by sequential actions of superoxide dismutase (SOD), and catalase converts superoxide into oxygen and water. Other elements in the cascade of ROS generation are small molecules derived from oxygen, which also include oxygen radicals [80].
Because ROS are biologically damaging, they need to be metabolized to prevent the damage they can cause when interacting with other compounds, for which the cell counts with mechanisms that avoid it like SOD. However, when the formation of ROS increases, they have the capacity to deteriorate mitochondrial function and jeopardize cell survival in different ways, where the mitochondrion seems to be responsible for regulating apoptosis [81]. ROS are a major threat to encode, transfer, and transport electrons and generate ATP by directly damaging mitochondrial DNA (mtDNA) which encodes 13 polypeptides, 12 transfer RNAs (tRNAs), and 2 ribosomal RNAs (rRNAs). All of them are essential in the chain of transport of electrons for the production of ATP, so when interacting with them, oxidative phosphorylation and therefore energy genesis is compromised [67]. ROS, and the release of proapoptotic proteins from the intermembrane space of mitochondria, triggers the activation of cell death.
The heart has the highest oxygen uptake rate in the human body, and the oxygen consumption is normally 8–13 mL 100 gˉ1 minˉ1 at rest [82]. The cellular sources in the genesis of ROS in the heart include cardiac myocytes, endothelial cells, and neutrophils. Within cardiac myocytes, ROS can be produced by several mechanisms, including the transport of mitochondrial electrons, NADPH oxidase (nicotinamide adenine dinucleotide phosphate oxidase), and xanthine dehydrogenase/xanthine oxidase. To meet the high demand for ATP synthesis, cardiac myocytes therefore have the highest volume density of mitochondria in the entire human body.
NADPH oxidase with its isoforms generically called NOX is the major source of ROS (reactive oxygen species) in biological systems. NOX proteins are involved in a plethora of pathophysiological conditions, so it is important to note that the functions of NOX proteins in different tissues are influenced by the activity of other oxidases and peroxidases, such as myeloperoxidase, xanthine oxidase, and hemoxygenase [83].
In the heart, the cardiomyocyte NADPH oxidase seems to be the main source of production of ROS from the heart in failure [84, 85].
NADPH oxidases are present in phagocytes and in a wide variety of non-phagocytic cells. NADPH generates superoxide by transferring electrons from NADPH into the cell through the membrane and coupling them to molecular oxygen to produce superoxide anion. Structurally, NADPH oxidase is an enzyme that has several components: it includes two integral membrane proteins, the glycoprotein gp. 1 Phox and the adapter protein p22 (phox), which together form the heterodimeric b558 flavocytochrome that form the nucleus of the enzyme. During the resting state, the multidomain regulatory subunits p40P (phox), p47 (phox), and p67 (Phox) are located in the cytosol organized as a complex. Activation of phagocytic NADPH oxidase occurs through a complex series of protein interactions.
The products that activate it are angiotensin II, endothelin-1, TNF-α, and mechanical forces. The cardiomyocyte NADPH oxidase and any other NADPH oxidase when stimulated generates large amounts of (O•2), which dismutes to H2O2; both in the tissue presence of iron and H2O2, increase the production of ROS, lead to the production of the HO• radical; these are highly reactive and can induce peroxidative damage of molecules within reach such as lipids, proteins, carbohydrates, nucleic acids, and membranes, resulting in the increase of reactive substances thiobarbituric acid (TBARS) in patients with heart failure.
This suggests that some pro-inflammatory products can activate a pathway to generate oxidative stress damage through the NADPH oxidase and increase the biological damage to the heart by ROS which correlates with left ventricular dysfunction [86]. Even more, the fact that NADPH oxidase is activated by pro-inflammatory products suggests a link with the genesis of oxidative stress.
Of the infectious processes in the heart on the balance of oxidants and antioxidants in the myocardium little is known. IE in which heart valves are usually affected, generating refractory congestive heart failure, is accompanied by a very important inflammatory response, both local and systemic with high circulating concentrations of IL-6, IL-2R, and IL-1β [87]. In the case of infective endocarditis, the interaction of the infectious agent and its products (chemotactic, formylated, and lipopolysaccharide peptides) with monocytes and polymorphonuclear cells can increase the production of ROS through the activation of NADPH oxidase, secondary to the inflammatory state.
IE induces an increase of pro-inflammatory cytokines, being able to stimulate ROS production in the myocardium and peroxidative damage to several molecules. The substances reactive to thiobarbituric acid (TBAR), in a study comparing cardiac tissue from patients with IE and patients with valvular heart disease (VHD) of rheumatic etiology; TBARs were increased 10 times more in IE than their controls with VHD [88].
In sepsis, endotoxins and cytokines stimulate the expression of inducible nitric oxide synthase (iNOS) and the overproduction of nitric oxide (NO) in various tissues; it also stimulates the excessive activity of NADPH oxidase that facilitates the expression of iNOS to produce large amounts of NO. The NADPH oxidases derived from ROS by activating the Jak2-IRF1 and JNK-AP1 pathways are necessary for the induction of iNOS. The main mechanism that regulates the activity of iNOS is the modulation of the transcription of the iNOS gene. The NO derived from iNOS and its metabolite peroxynitrite can contribute to the pathological alterations observed in sepsis, such as endothelial dysfunction, hypotension, and multiple organ failure [89].
The peroxynitrite anion ONOO−
The composition of metabolites such as amino acids, intermediate products of the Krebs cycle, and acylcarnitines (metabolome) and protein complement expressed in cells, tissues, or body fluids (proteome) of survivors of sepsis and non-survivors was analyzed in patients who studied with sepsis by three different pathogens, S. pneumoniae, S. aureus, or E. coli. The main differences between survivors and non-survivors were those highlighted in their metabolome and proteome. For example, nine proteins involved in the transport of fatty acids were decreased in non-survivors of sepsis, suggesting a defect in β-oxidation. The nonacceptance and nonuse of fatty acids by the mitochondria led to an accumulation of acylcarnitines in the plasma; another predictive marker is that glycolysis and gluconeogenesis were also markedly different. Survivors of sepsis showed decreased levels of citrate, malate, glycerol, glycerol 3-phosphate, phosphate, and glucogenic and ketogenic amino acids, while non-survivors showed elevated levels of citrate, malate, pyruvate, dihydroxyacetone, lactate, phosphate, and gluconeogenic amino acids [90]. That is to say that the pathways for the transport of fatty acids, as well as glycolysis and gluconeogenesis, are damaged, so the substrate is low, and they are not used by the mitochondria.
Acetylome analysis identified a subpopulation of mitochondrial proteins that was sensitive to changes in the NADH/NAD+ ratio. Hyperacetylation induced by mitochondrial dysfunction is a positive regulator of pathological remodeling in the heart of mice with primary or acquired mitochondrial dysfunction, as well as in humans with heart failure. Hyperacetylation of mitochondrial malate–aspartate shuttle (MAS) proteins impaired the transport and oxidation of cytosolic NADH in the mitochondria, resulting in altered cytosolic redox state and energy deficiency. Furthermore, acetylation of oligomycin-sensitive conferring protein at lysine-70 in adenosine triphosphate synthase complex promoted its interaction with cyclophilin D and sensitized the opening of mitochondrial permeability transition pore. There are two different mechanisms that point to the proteins of hyperacetylation, i.e., MAS and the regulators of mitochondrial permeability transition pore (mPTP), which mediate an increase in heart failure. Both could be fixed by normalizing the NAD+ redox balance either genetically or pharmacologically [91].
Q and cytochrome c (Cytc) are mobile. Q diffuses rapidly within the membrane, while cytochrome c is a soluble protein that contains a peptide sequence located at the C-terminus of the protein [92] that allows it to cross the cell membranes in a nontraditional way. This property of Cytc was used in a study in mice, which were subject to ligation and cecal puncture; they underwent sepsis and damage to mitochondrial respiration, which was restored with the injection i.v. of Cytc [93]. The treatment led to an uptake of Cytc into the cardiomyocytes, and survival increased from 15% for the sepsis control group to about 50% in mice that were also injected with Cytc [94].
The death of cells of the immune system by deregulated apoptosis contributes to the dysfunction of the immune system and multiple organ failure (MOF) which is observed in sepsis. The immune cells most affected by this dysregulated apoptotic cell death appear to be lymphocytes [95]. Extensive lymphocytic apoptosis mediated by caspase-3 in sepsis may contribute to impaired immune response in septic patients [96]. Lymphocyte loss occurs by both death receptor and mitochondrial-mediated apoptosis, suggesting that there may be multiple triggers for lymphocyte apoptosis [97, 98].
Apoptosis in the immune system is a pathological event in sepsis which has been considered a therapeutic goal. Studies on sepsis in experimental animals suggest that the loss of lymphocytes during sepsis may be due to deregulated apoptosis and that it appears to be secondary to a variety of mediators that carry out both “intrinsic” and “extrinsic” cell death pathways.
In experimental animals, lymphocyte apoptosis is frequently seen 12 h after the onset of experimental polymicrobial sepsis in the thymus, spleen, and lymphoid tissues associated with the intestine. It has been suggested that deregulated lymphocytic apoptosis results in reduced septic survival through loss of lymphocytes, resulting in multiple organ failure and ultimately death. Lymphocyte apoptosis in the thymus appears to occur 4 h after the onset of sepsis and is independent of the effects of endotoxin or death receptors. Apoptosis in the spleen appears to be particularly important in mortality from sepsis, by an increase of the splenic apoptosis of lymphocytes in experimental animals after the cecal ligation and puncture (CLP) which results in a reduced survival [99].
In septic humans apoptosis does not seem to be generalized, since in these patients only extensive lymphocytic apoptosis was demonstrated, which suggests a damaged immune response, suggesting that other mechanisms apart from cell death participate in the conditions associated with mortality [100]. For example, hyperglycemia induces the expression of leukocyte adhesion molecules, such as the intercellular adhesion molecule (ICAM) and vascular cell adhesion molecule (VCAM), which is suppressed by treatment with insulin. Another example is the impairment induced by hyperglycemia in the function of neutrophils, including chemotaxis, phagocytosis, and respiratory function, which is attenuated with insulin [101].
As we observed, the epidemiology of IE has changed over time. S. aureus is currently the most important pathological agent as a cause of IE [4, 102]. The age group with greater participation is the older adult due to their comorbidities, especially cardiac ones, with the need for valve prosthesis placement, and vascular approach for the placement of cardiac pacemakers.
The existence of an immunogenetic influence in the risk and outcomes of infectious diseases has been well stablished. In the cases of IE and sepsis, investigation is ongoing to clearly define the specific genetic anomalies that contribute to this influence. The study of SNPs has been a good start in the understanding of the phenomena; nevertheless at the light of the information derived from their study, they do not seem sufficient to explain the whole participation of genetics in the sepsis and IE equation. Other types of genetic abnormalities might also participate, and it might be worth exploring [103]. Even though there is a large body of studies with positive results, there are also lots of contradictory and conflicting findings that make it difficult to make definitive conclusions. Even more, according to a systematic review made to determine the methodological quality of SNP association studies with sepsis, most of the studies could improve a lot methodologically speaking in terms of control group selection, genetic assay technique, study blinding, statistical interpretation, study replication, study size, and power.
Finally, the sequence of events that begin with an infectious state, such as IE, alerts and promotes inflammation through the immune system, both cellular and humoral to eliminate the infectious agent; however, this has the ability to evade the immune system.
In its evolution, the germ also generated the possibility of survival through the acquisition of resistance to external agents, such as antibiotics, which can perpetuate the septic process, increasing the production of reactive O2 species both locally (cell-mitochondria) and systemic level (neutrophil-monocytes-macrophage-endothelium) together with the products that generate the interaction infectious agent-immune system.
The activity of antioxidant enzymes is exceeded, so that ROS cannot be eliminated, generating a state of oxidative stress, with a profound effect on the mitochondrial level by breaking the chain of electron transport, and, consequently, the genesis of the energy is compromised.
The repercussion of this sequence of events, both at the cardiac level and at the systemic level, is manifested by the failure of one or several organs.
In a schematic way, the sequence of events of a patient with IE who has a severe evolution and finally dies of multiple organ failure is shown (Figure 3).
Schematic representation of the sequence of events of a patient with IE who has a severe evolution and finally dies of multiple organ failure.
Different studies explore areas of compromise such as metabolome and proteome in which it is observed that glycolysis, gluconeogenesis, and fatty acid transport are damaged, so the substrate is low and the few substrates are not used by the mitochondria, which generates attention in processes to be repaired.
In another (acetylome) the possibility of normalizing the NAD + redox balance is observed both genetically and pharmacologically in the treatment of heart failure [91].
The observations of the behavior of cytochrome c, being a mobile complex molecule and crossing cell membranes, made it possible for cytochrome c to enter into cardiomyocytes to improve mitochondrial respiration, improving the survival of septic mice [92, 93, 94]. This open a very attractive opportunity in the treatment of septic patients with heart failure as in IE when in the future we use complex molecules, i.v., in the treatment of these patients.
There are still many areas in which it is necessary to continue researching in the clinical area as well as in the bacteriological, biochemical, and biomolecular areas in addition to other types of tools to observe systemic inflammation, through mathematical modulation and systems-based models of inflammation [104, 105], and the severity of a septic patient due to the complexity of losing the cardiac bioelectrical signal and how it recovers the complexity if the patient survives the septic event have also been considered [106].
Vet. Laura Yavarik Alvarado Avila. Faculty of Veterinary Medicine. National Autonomous University of Mexico UNAM, Mexico City, Mexico. We appreciate the support in the corrections and spelling suggestions made in this chapter on Infective Endocarditis.
None.
MRSA | methicillin-resistant Staphylococcus aureus |
HssRS | hem sensor system |
hrtAB | hem regulator transporter |
bNOS | nitric oxide synthase hemoprotein |
TNF | tumor necrosis factor alpha |
IL | interleukin |
HMGB1 | high-mobility group box 1 protein |
GRO alpha | growth-regulated oncogene |
RANTES | regulated upon activation, normal T-cell expressed, and secreted |
EE | endocardial endothelium |
Et-1 | endothelin-1 |
ON | nitric oxide |
PG | prostaglandins |
VEGF | vascular endothelial growth factor |
SNP | single-nucleotide polymorphism |
GWAS | genome-wide association studies |
LPS | lipopolysaccharides |
A | adenosine |
G | guanine |
PBMC | peripheral blood mononuclear cells |
IL-RNa* | The interleukin 1receptor antagonist gene |
APACHE II score | acute physiology and chronic health evaluation |
SIRS | systemic inflammatory response syndrome |
ICU | intensive care unit |
TLR | toll-like receptor |
MyD88 | adaptor-like protein |
FER gene | tyrosine-protein kinase |
TREMs | triggering receptor expressed on myeloid cells |
ADP | adenosine diphosphate |
NADH | nicotinamide adenine dinucleotide phosphate |
FADH | flavin adenine dinucleotide |
ROS | reactive oxygen species |
IMM | inner mitochondrial membrane |
OH• | hydroxyl |
RO•2 | peroxyl |
RO• | alkoxyl |
HOCl | hypochlorous acid |
O3 | ozone |
½O2 | singlet oxygen |
H2O2 | hydrogen peroxide |
RNS | reactive nitrogen species |
SOD | superoxide dismutases |
2O•2 | superoxide free radical |
CuZnSOD | copper, zinc-superoxide dismutase |
mtDNA | mitochondrial DNA |
tRNAs | transfer RNAs |
rRNAs | ribosomal RNAs |
NOX | NADPH oxidase generically called |
phox | adapter protein p22 |
TBARS | reactive substances thiobarbituric acid |
iNOS | inducible nitric oxide synthase |
NO | nitric oxide |
JNK-AP1 | Jak2-IRF1 pathway genes (IFNGR1, IFNGR2, JAK1, JAK2, STAT1, IRF1) |
MOF | multiple organ failure |
MODS | multi-organ dysfunction syndrome |
CLP | cecal ligation and puncture |
ICAM | intercellular adhesion molecule |
VCAM | vascular cell adhesion molecule |
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