Academic literature on the topic 'India Telengana region'

Site specific geoid model is prerequisite for accurate determination of orthometric heights. No geoid model has been developed so far for India or any of its part. So, development of a geoid model for India or its part is of utmost need to make use of GNSS data towards determination of orthometric heights. In this research work, an attempt has been made to develop geoid undulation models by gravimetric method using Molodensky’s concept. Component parameters in line with the Remove – Compute – Restore (RCR) technique have been used recursively. Models have been developed for two study areas: one of these lies in and around Dehradun (30° 19′ N, 75° 04′E) in Uttarakhand state, India in lower Himalayan region having highly undulating topography and the other near Hyderabad (17° 30′N, 78°30′E) in Telengana state of India having gentle topography. The model has been tested for 7 stations in the first study area and accuracy has been found to be 17.5 cm; whereas, for the second area accuracy has been found to be 7.0 cm for 24 test stations. Further, the performances of the developed models have been evaluated with those from three global geoid models namely EIGEN6C4, EIGEN6C3stat and EGM2008; and have been found to be similar or better in case of first study and for second study area far more superior. Thus, local/regional geoid undulation model requiring accuracy better than 20 cm for any study area may be developed adopting the method. However, the optimality in the number and density of gravity stations may be considered as a future scope of work.

In mid autumn 1992 the South American leaf-miner Liriomyza huidobrensis (Blanchard) was firstly recorded on greenhouse cucumbers, melons, beans and outdoor beans, broad beans and lettuce in the following locations of Crete: 1) Mires, Tymbaki, Antiskari at Messara valley of Southern Crete 2) Filissia in the midland and 3) Platanias and Kokini Chani in Northern Crete. The following year L. huidobrensis had spread all over Crete (Ierapetra, Stomion, Koutsoura, Chania etc.) while melon and potato leaves which were sent to our laboratory from mainland Greece, Pirgos (Peloponissos) and Chalkida (Evia island), were found heavily damaged by the same leafmincr. The different types of mines (it usually mines the leaf alongside the veins), the colour of pupae (blackish) and the sudden attack of some plants (lettuce, broad beans, onions) which have never been damaged by the known local leafminers as yet, indicated that it was a newly introduced species. The new leafminer alerted the growers due to the high crop losses on outdoor and greenhouse vegetables (lettuce, broad beans, beans etc) despite the frequent application of registered insecticides. In the following year an extensive survey started to investigate its distribution and host plant in the greenhouses and outdoor vegetables and ornamentals in Crete. L. huidobrensis, a quarantine insect, is a polyphagous leafminer distributed in most biogeographical regions. It is native in South American countries from which it was spread soon in North America, Asia and Europe. In Europe it was first noticed in Holland, England and France in 1989 causing considerable damage to vegetables and ornamentals. It mines the spongy mesophyll reducing the photosynthesis more than the other leafminers. This type of mines is not easily visible, unless the leaf is observed through transmitted sunlight or artificial light. This method was applied to ensure accurate detection of larvae and their parasitoids under a stereoscope. In our observations we found mines in leaves and cucumber fruits as well. Studies of its life cycle on greenhouse beans and melons revealed that most pupae (98%) remain on the bean leaves outside a hole in the autumn, while a few fall on the ground. The number of pupae collected from ten leaves per 24h was: 5.7±1.6 on lettuce, 17.07±4.1 on beans, 24.2±7.3 on melon, 6.0±2 on broad bean leaves. During the survey the following host plants were recorded: Chemical control could be effective by applying certain insecticides (abamectin, triazophos, imidacloprid, heptenophos etc.) while biological control seems to be rather effective by the known parasitoids, which are already used against the tomato leafminer Liriomyza bryoniae Kalt. Biological control of L. huidobrensis was effective on lettuce by means of repeated releases of Dacnusa sibirica Telenga and Diglyphus isaea (Walker) but so far native parasitoids proved to be able to control the pest. Mass trapping with coloured sticky traps seems to be also a potential method in IPM programmes. The mortality of pupae collected from heavily treated greenhouse plants with insecticides varied from 65 to 74% while that of untreated plants was between 18-25%. No pupal parasitoids were recorded but D. isaea and D. sibirica were both recorded as larval parasitoids. D. isaea was abundant all over the year while D. sibirica only in spring-summer period. These two parasitoids were able, in certain cases, to control sufficiently the leaf miner in untreated greenhouse cucumber and bean plants. The neem seed extract (Azadirachta indica) applied on outdoor tomatoes in Antiskari (Southern Crete) proved harmless both on hymenoptera and on the mirid predator Macrolophus caliginogus Wagner. Despite its weak larvicide action on L. huidobrensis it was very effective in conjuction with parasitoids consisting of a promising candidate in IPM programmes.

A checklist of world species of Microgastrinae parasitoid wasps (Hymenoptera: Braconidae) is provided. A total of 81 genera and 2,999 extant species are recognized as valid, including 36 nominal species that are currently considered as species inquirendae. Two genera are synonymized under Apanteles. Nine lectotypes are designated. A total of 318 new combinations, three new replacement names, three species name amendments, and seven species status revised are proposed. Additionally, three species names are treated as nomina dubia, and 52 species names are considered as unavailable names (including 14 as nomina nuda). A total of three extinct genera and 12 extinct species are also listed. Unlike in many previous treatments of the subfamily, tribal concepts are judged to be inadequate, so genera are listed alphabetically. Brief diagnoses of all Microgastrinae genera, as understood in this paper, are presented. Illustrations of all extant genera (at least one species per genus, usually more) are included to showcase morphological diversity. Primary types of Microgastrinae are deposited in 108 institutions worldwide, although 76% are concentrated in 17 collections. Localities of primary types, in 138 countries, are reported. Recorded species distributions are listed by biogeographical region and by country. Microgastrine wasps are recorded from all continents except Antarctica; specimens can be found in all major terrestrial ecosystems, from 82°N to 55°S, and from sea level up to at least 4,500 m a.s.l. The Oriental (46) and Neotropical (43) regions have the largest number of genera recorded, whereas the Palaearctic region (28) is the least diverse. Currently, the highest species richness is in the Palearctic region (827), due to more historical study there, followed by the Neotropical (768) and Oriental (752) regions, which are expected to be the most species rich. Based on ratios of Lepidoptera and Microgastrinae species from several areas, the actual world diversity of Microgastrinae is expected to be between 30,000–50,000 species; although these ratios were mostly based on data from temperate areas and thus must be treated with caution, the single tropical area included had a similar ratio to the temperate ones. Almost 45,000 specimens of Microgastrinae from 67 different genera (83% of microgastrine genera) have complete or partial DNA barcode sequences deposited in the Barcode of Life Data System; the DNA barcodes represent 3,545 putative species or Barcode Index Numbers (BINs), as estimated from the molecular data. Information on the number of sequences and BINs per genus are detailed in the checklist. Microgastrinae hosts are here considered to be restricted to Eulepidoptera, i.e., most of the Lepidoptera except for the four most basal superfamilies (Micropterigoidea, Eriocranioidea, Hepialoidea and Nepticuloidea), with all previous literature records of other insect orders and those primitive Lepidoptera lineages being considered incorrect. The following nomenclatural acts are proposed: 1) Two genera are synonymyzed under Apanteles: Cecidobracon Kieffer & Jörgensen, 1910, new synonym and Holcapanteles Cameron, 1905, new synonym; 2) Nine lectotype designations are made for Alphomelon disputabile (Ashmead, 1900), Alphomelon nigriceps (Ashmead, 1900), Cotesia salebrosa (Marshall, 1885), Diolcogaster xanthaspis (Ashmead, 1900), Dolichogenidea ononidis (Marshall, 1889), Glyptapanteles acraeae (Wilkinson, 1932), Glyptapanteles guyanensis (Cameron, 1911), Glyptapanteles militaris (Walsh, 1861), and Pseudapanteles annulicornis Ashmead, 1900; 3) Three new replacement names are a) Diolcogaster aurangabadensis Fernandez-Triana, replacing Diolcogaster indicus (Rao & Chalikwar, 1970) [nec Diolcogaster indicus (Wilkinson, 1927)], b) Dolichogenidea incystatae Fernandez-Triana, replacing Dolichogenidea lobesia Liu & Chen, 2019 [nec Dolichogenidea lobesia Fagan-Jeffries & Austin, 2019], and c) Microplitis vitobiasi Fernandez-Triana, replacing Microplitis variicolor Tobias, 1964 [nec Microplitis varicolor Viereck, 1917]; 4) Three names amended are Apanteles irenecarrilloae Fernandez-Triana, 2014, Cotesia ayerzai (Brèthes, 1920), and Cotesia riverai (Porter, 1916); 5) Seven species have their status revised: Cotesia arctica (Thomson, 1895), Cotesia okamotoi (Watanabe, 1921), Cotesia ukrainica (Tobias, 1986), Dolichogenidea appellator (Telenga, 1949), Dolichogenidea murinanae (Capek & Zwölfer, 1957), Hypomicrogaster acarnas Nixon, 1965, and Nyereria nigricoxis (Wilkinson, 1932); 6) New combinations are given for 318 species: Alloplitis congensis, Alloplitis detractus, Apanteles asphondyliae, Apanteles braziliensis, Apanteles sulciscutis, Choeras aper, Choeras apollion, Choeras daphne, Choeras fomes, Choeras gerontius, Choeras helle, Choeras irates, Choeras libanius, Choeras longiterebrus, Choeras loretta, Choeras recusans, Choeras sordidus, Choeras stenoterga, Choeras superbus, Choeras sylleptae, Choeras vacillatrix, Choeras vacillatropsis, Choeras venilia, Cotesia asavari, Cotesia bactriana, Cotesia bambeytripla, Cotesia berberidis, Cotesia bhairavi, Cotesia biezankoi, Cotesia bifida, Cotesia caligophagus, Cotesia cheesmanae, Cotesia compressithorax, Cotesia delphinensis, Cotesia effrena, Cotesia euphobetri, Cotesia elaeodes, Cotesia endii, Cotesia euthaliae, Cotesia exelastisae, Cotesia hiberniae, Cotesia hyperion, Cotesia hypopygialis, Cotesia hypsipylae, Cotesia jujubae, Cotesia lesbiae, Cotesia levigaster, Cotesia lizeri, Cotesia malevola, Cotesia malshri, Cotesia menezesi, Cotesia muzaffarensis, Cotesia neptisis, Cotesia nycteus, Cotesia oeceticola, Cotesia oppidicola, Cotesia opsiphanis, Cotesia pachkuriae, Cotesia paludicolae, Cotesia parbhanii, Cotesia parvicornis, Cotesia pratapae, Cotesia prozorovi, Cotesia pterophoriphagus, Cotesia radiarytensis, Cotesia rangii, Cotesia riverai, Cotesia ruficoxis, Cotesia senegalensis, Cotesia seyali, Cotesia sphenarchi, Cotesia sphingivora, Cotesia transuta, Cotesia turkestanica, Diolcogaster abengouroui, Diolcogaster agama, Diolcogaster ambositrensis, Diolcogaster anandra, Diolcogaster annulata, Diolcogaster bambeyi, Diolcogaster bicolorina, Diolcogaster cariniger, Diolcogaster cincticornis, Diolcogaster cingulata, Diolcogaster coronata, Diolcogaster coxalis, Diolcogaster dipika, Diolcogaster earina, Diolcogaster epectina, Diolcogaster epectinopsis, Diolcogaster grangeri, Diolcogaster heterocera, Diolcogaster homocera, Diolcogaster indica, Diolcogaster insularis, Diolcogaster kivuana, Diolcogaster mediosulcata, Diolcogaster megaulax, Diolcogaster neglecta, Diolcogaster nigromacula, Diolcogaster palpicolor, Diolcogaster persimilis, Diolcogaster plecopterae, Diolcogaster plutocongoensis, Diolcogaster psilocnema, Diolcogaster rufithorax, Diolcogaster semirufa, Diolcogaster seyrigi, Diolcogaster subtorquata, Diolcogaster sulcata, Diolcogaster torquatiger, Diolcogaster tristiculus, Diolcogaster turneri, Diolcogaster vulcana, Diolcogaster wittei, Distatrix anthedon, Distatrix cerales, Distatrix cuspidalis, Distatrix euproctidis, Distatrix flava, Distatrix geometrivora, Distatrix maia, Distatrix tookei, Distatrix termina, Distatrix simulissima, Dolichogenidea agamedes, Dolichogenidea aluella, Dolichogenidea argiope, Dolichogenidea atreus, Dolichogenidea bakeri, Dolichogenidea basiflava, Dolichogenidea bersa, Dolichogenidea biplagae, Dolichogenidea bisulcata, Dolichogenidea catonix, Dolichogenidea chrysis, Dolichogenidea coffea, Dolichogenidea coretas, Dolichogenidea cyane, Dolichogenidea diaphantus, Dolichogenidea diparopsidis, Dolichogenidea dryas, Dolichogenidea earterus, Dolichogenidea ensiger, Dolichogenidea eros, Dolichogenidea evadne, Dolichogenidea falcator, Dolichogenidea gelechiidivoris, Dolichogenidea gobica, Dolichogenidea hyalinis, Dolichogenidea iriarte, Dolichogenidea lakhaensis, Dolichogenidea lampe, Dolichogenidea laspeyresiella, Dolichogenidea latistigma, Dolichogenidea lebene, Dolichogenidea lucidinervis, Dolichogenidea malacosomae, Dolichogenidea maro, Dolichogenidea mendosae, Dolichogenidea monticola, Dolichogenidea nigra, Dolichogenidea olivierellae, Dolichogenidea parallelis, Dolichogenidea pelopea, Dolichogenidea pelops, Dolichogenidea phaenna, Dolichogenidea pisenor, Dolichogenidea roepkei, Dolichogenidea scabra, Dolichogenidea statius, Dolichogenidea stenotelas, Dolichogenidea striata, Dolichogenidea wittei, Exoryza asotae, Exoryza belippicola, Exoryza hylas, Exoryza megagaster, Exoryza oryzae, Glyptapanteles aggestus, Glyptapanteles agynus, Glyptapanteles aithos, Glyptapanteles amenophis, Glyptapanteles antarctiae, Glyptapanteles anubis, Glyptapanteles arginae, Glyptapanteles argus, Glyptapanteles atylana, Glyptapanteles badgleyi, Glyptapanteles bataviensis, Glyptapanteles bistonis, Glyptapanteles borocerae, Glyptapanteles cacao, Glyptapanteles cadei, Glyptapanteles cinyras, Glyptapanteles eryphanidis, Glyptapanteles euproctisiphagus, Glyptapanteles eutelus, Glyptapanteles fabiae, Glyptapanteles fulvigaster, Glyptapanteles fuscinervis, Glyptapanteles gahinga, Glyptapanteles globatus, Glyptapanteles glyphodes, Glyptapanteles guierae, Glyptapanteles horus, Glyptapanteles intricatus, Glyptapanteles lamprosemae, Glyptapanteles lefevrei, Glyptapanteles leucotretae, Glyptapanteles lissopleurus, Glyptapanteles madecassus, Glyptapanteles marquesi, Glyptapanteles melanotus, Glyptapanteles melissus, Glyptapanteles merope, Glyptapanteles naromae, Glyptapanteles nepitae, Glyptapanteles nigrescens, Glyptapanteles ninus, Glyptapanteles nkuli, Glyptapanteles parasundanus, Glyptapanteles penelope, Glyptapanteles penthocratus, Glyptapanteles philippinensis, Glyptapanteles philocampus, Glyptapanteles phoebe, Glyptapanteles phytometraduplus, Glyptapanteles propylae, Glyptapanteles puera, Glyptapanteles seydeli, Glyptapanteles siderion, Glyptapanteles simus, Glyptapanteles speciosissimus, Glyptapanteles spilosomae, Glyptapanteles subpunctatus, Glyptapanteles thespis, Glyptapanteles thoseae, Glyptapanteles venustus, Glyptapanteles wilkinsoni, Hypomicrogaster samarshalli, Iconella cajani, Iconella detrectans, Iconella jason, Iconella lynceus, Iconella pyrene, Iconella tedanius, Illidops azamgarhensis, Illidops lamprosemae, Illidops trabea, Keylimepie striatus, Microplitis adisurae, Microplitis mexicanus, Neoclarkinella ariadne, Neoclarkinella curvinervus, Neoclarkinella sundana, Nyereria ituriensis, Nyereria nioro, Nyereria proagynus, Nyereria taoi, Nyereria vallatae, Parapanteles aethiopicus, Parapanteles alternatus, Parapanteles aso, Parapanteles atellae, Parapanteles bagicha, Parapanteles cleo, Parapanteles cyclorhaphus, Parapanteles demades, Parapanteles endymion, Parapanteles epiplemicidus, Parapanteles expulsus, Parapanteles fallax, Parapanteles folia, Parapanteles furax, Parapanteles hemitheae, Parapanteles hyposidrae, Parapanteles indicus, Parapanteles javensis, Parapanteles jhaverii, Parapanteles maculipalpis, Parapanteles maynei, Parapanteles neocajani, Parapanteles neohyblaeae, Parapanteles nydia, Parapanteles prosper, Parapanteles prosymna, Parapanteles punctatissimus, Parapanteles regalis, Parapanteles sarpedon, Parapanteles sartamus, Parapanteles scultena, Parapanteles transvaalensis, Parapanteles turri, Parapanteles xanthopholis, Pholetesor acutus, Pholetesor brevivalvatus, Pholetesor extentus, Pholetesor ingenuoides, Pholetesor kuwayamai, Promicrogaster apidanus, Promicrogaster briareus, Promicrogaster conopiae, Promicrogaster emesa, Promicrogaster grandicula, Promicrogaster orsedice, Promicrogaster repleta, Promicrogaster typhon, Sathon bekilyensis, Sathon flavofacialis, Sathon laurae, Sathon mikeno, Sathon ruandanus, Sathon rufotestaceus, Venanides astydamia, Venanides demeter, Venanides parmula, and Venanides symmysta.

India is consisting of 29 states and 7 union territories, including a national capital, Delhi. Elevated concentrations (>10 g l ) of arsenic (As) in ground water (GW)  -1 of many states of India have become a major concern in recent years. Up to now about 0.2 million GW samples have been analyzed for As contamination from all over India by various researchers and Government agencies. About 90% of these cover only the Eastern part of India while several states and UTs are still unexplored. However, from the available data, GW of eighteen Indian states and three union territories has been found to be As contaminated to different extents through natural or anthropogenic origin. Among these, As >300 μg l has been reported from at least one locality from fourteen states. The -1 maximum level of As (7350 μg l ) in GW has been reported from a highly industrialized -1 area, Patancheru in Medak district of Andhra Pradesh. However, the gravity of problem is more in West Bengal followed by Bihar and Uttar Pradesh. Five out of eight North-Eastern states are also affected by As contamination. Manipur is ranked first and Assam as second followed by Arunachal Pradesh, Tripura and Nagaland. The GW in these regions is naturally As enriched, and therefore wide spatial distribution of As has been found in these areas. In North India, Punjab and Haryana and in South India, Andhra Pradesh and Karnataka are suffering with GW As contamination. Low level of As (up to 17 μg l ) has also -1 been reported in Tamil Nadu from South India. Many of the states like Jammu and Kashmir, Uttarakhand, Odisha, Gujrat, Kerala, Telengana, Goa etc. are still unexplored for GW As contamination. Thus, according to current reports out of 640 districts in India, 141 are As affected (As >10 g l-1), among them 120 are above 50 g l-1. Considering its severity, the issue of As contamination in drinking water has been taken up by the Government of India and mitigation efforts are being initiated. In order to provide safe drinking water, different agencies/ organizations have developed eco-friendly, cost effective devices/ filtration techniques having higher As removal capacity. Here we elucidated the current status of GWAs contamination in different states of India and the new developments of mitigation options.