# Medicinal Plants for the Treatment of Local Tissue Damage Induced by Snake Venoms: An Overview from Traditional Use to Pharmacological Evidence

Authors: Juliana Félix-Silva; Arnóbio Antônio Silva-Junior; Silvana Maria Zucolotto; Matheus de Freitas Fernandes-Pedrosa
Journal: Evidence-Based Complementary and Alternative Medicine
 (2017)

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## Abstract

Snakebites are a serious problem in public health due to their high morbimortality. Most of snake venoms produce intense local tissue damage, which could lead to temporary or permanent disability in victims. The available specific treatment is the antivenom serum therapy, whose effectiveness is reduced against these effects. Thus, the search for complementary alternatives for snakebite treatment is relevant. There are several reports of the popular use of medicinal plants against snakebites worldwide. In recent years, many studies have been published giving pharmacological evidence of benefits of several vegetal species against local effects induced by a broad range of snake venoms, including inhibitory potential against hyaluronidase, phospholipase, proteolytic, hemorrhagic, myotoxic, and edematogenic activities. In this context, this review aimed to provide an updated overview of medicinal plants used popularly as antiophidic agents and discuss the main species with pharmacological studies supporting the uses, with emphasis on plants inhibiting local effects of snake envenomation. The present review provides an updated scenario and insights into future research aiming at validation of medicinal plants as antiophidic agents and strengthens the potentiality of ethnopharmacology as a tool for design of potent inhibitors and/or development of herbal medicines against venom toxins, especially local tissue damage.


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## Body

## 1. Introduction

Snakebites are a serious public health problem in many regions around the world, particularly in Africa, Asia, Latin America, and parts of Oceania [1]. Conservative data indicate that, worldwide, there are between 1.2 and 5.5 million snakebites every year, leading to 25,000 to 125,000 deaths [2]. Despite its significant impact on human health, this condition remains largely neglected by national and international health authorities, funding agencies, pharmaceutical companies, patients’ organizations, and health advocacy groups [1]. Thus, snake envenomation is included since 2009 in World Health Organization (WHO) list of Neglected Tropical Diseases (NTDs) [3]. Envenoming and deaths resulting from snakebites are a particularly important public health problem in the rural tropics. Populations in these regions experience high morbidity and mortality because of poor access to health services, which are often suboptimal, as well as other NTDs, which are associated with poverty [3, 4].Snakes with major clinical importance belong to the families Elapidae (African and Asian cobras, Asian kraits, African mambas, American coral snakes, Australian and New Guinean venomous snakes, and sea snakes) and Viperidae (Old World vipers, American rattlesnakes and pit vipers, and Asian pit vipers) [5]. After production, snake venom is injected in the victim via tubular or channeled fangs [6]. Biochemically, venoms are complex mixtures of pharmacologically active proteins and polypeptides, acting in concert to help in immobilizing the prey [7]. The most common toxins in snake venoms are snake venom metalloproteinases (SVMPs), phospholipases A2 (PLA2s), snake venom serine proteinases (SVSPs), acetylcholinesterase (AChE), L-amino acid oxidases (LAAOs), nucleotidases, and snake venom hyaluronidases (SVHs) [7].Biological properties of snake venom components are peculiar to each species, but in general, the main clinical effects of snake envenomation are immediate and prominent local tissue damage (including myonecrosis, dermonecrosis, hemorrhage, and edema), coagulation disorders (consumption coagulopathy and spontaneous systemic bleeding), cardiovascular alterations (hypotension, hypovolemic shock, and myocardial damage), renal alterations (which could evolve into acute kidney injure), neurotoxic action (descending paralysis, progressing from ptosis and external ophthalmoplegia to bulbar, respiratory muscle, and total flaccid paralysis), generalized rhabdomyolysis with myoglobinuria, and intravascular haemolysis [5, 8].The only available specific treatment is the antivenom serum therapy, which consists of a pool of neutralizing immunoglobulins, or immunoglobulin fragments, purified from the plasma of animals hyperimmunized against snake venoms or specific toxins. Its effectiveness consists in its ability to provide to the patient antibodies with a high affinity to snake venom, aiming to eliminate the toxins responsible for toxicity of the envenoming, mitigating the progress of toxic effects induced by snake venom components [9]. However, the antivenom has some limitations, such as poor ability to treat local effects, risk of immunological reactions, high cost, and difficult access in some regions [8–10]. If antivenom administration is initiated rapidly after envenomation, neutralization of systemic effects is usually achieved successfully; however, neutralization of local tissue damage is more difficult [8]. Furthermore, the availability and accessibility of antivenoms is limited in many regions, such as Sub-Saharan Africa, Asia, and, to a lesser extent, Latin America, which could aggravate even more this picture [1]. Thus, this inability to treat local effects, as well as the increased time between accident and treatment, is the main reason for the temporary or permanent disability observed in many victims, which can lead to serious social, economic, and health negative impacts, given that most victims live in rural areas [3].In this context, the search for complementary therapies to treat snakebites is relevant and medicinal plants could be highlighted as a rich source of natural inhibitors and pharmacologically active compounds [6, 11–13]. There are several reports of the popular use of medicinal plants against snakebites around the world, especially in tropical and subtropical regions such as Asia, Africa, and South America [14, 15]. The rural and tribal people living in remote areas greatly depend on folk medicines for the treatment of bites from any venomous creatures [16]. The use of medicinal plants against snakebites is a historical practice throughout the human history, and this knowledge has been transferred among the rural communities from generation after generation [17]. Nowadays, these herbal antidotes used in folk traditional medicine gained much attention by toxinologists worldwide as a tool for design of potent inhibitors against snake venom toxins. The potential advantages of antiophidic plants are their possible low cost, easy access, stability at room temperature, and ability to neutralize a broad spectrum of toxins, including the local tissue damage [12, 15–17].So, the objective of this review is to provide an updated overview of medicinal plants used popularly as antiophidic and discuss the main species with pharmacological studies supporting the uses, with emphasis on plants inhibiting local effects of snake envenomation, since this is a critical effect of snake venoms that could lead to relevant sequel to victims. A review of the main botanical families popularly used as antiophidic is presented, including the main species and forms of popular use of them. Then, studies supporting their popular use are discussed, as well as the advantages of this kind of approach for treatment of snake venom accident.


## 2. Methodology

An extensive review of the literature was undertaken in different scientific sources, such as PubMed (https://www.ncbi.nlm.nih.gov/pubmed), Science Direct (http://www.sciencedirect.com/), Scopus (https://www.scopus.com/), Web of Science (http://www.webofknowledge.com/),“Literatura Latino-Americana e do Caribe em Ciências da Saúde” (LILACS) (http://lilacs.bvsalud.org/), Scientific Electronic Library Online (SciELO) (http://www.scielo.org/), Google Scholar (https://scholar.google.com.br/), Cochrane Library (http://www.cochranelibrary.com/), and Centre for Reviews and Dissemination (CRD) (http://www.crd.york.ac.uk/CRDWeb).The study database included original articles published in peer-reviewed journals, as well as books, thesis, dissertations, patents, and other reports covering antiophidic plants (ethnopharmacological surveys, original articles, or reviews), dated until December 2016. For the online search, where applicable, the following search strategy was employed: (“plant” OR “plants” OR “plant extract” OR “vegetal” OR “vegetal species” OR “vegetal extract” OR “traditional medicine” OR “alternative medicine” OR “complementary therapy” OR “natural medicine” OR “ethnopharmacology” OR “ethnobotany” OR “herbal medicine” OR “herb” OR “herbs” OR “decoction” OR “tea” OR “infusion” OR “macerate”) AND (“snake venom” OR “snake” OR “snakes” OR “snakebite” OR “snakebites” OR “antivenom” OR “antivenoms” OR “anti-venom” OR “anti-venoms” OR “antivenin” OR “antivenins” OR “anti-venin” OR “anti-venins” OR “antiophidian” OR “antiophidic” OR “snake envenomation” OR “antitoxin” OR “antitoxins” OR “snake antidote” OR “snake antidotes” OR “snake venom neutralization” OR “snake venom inhibition” OR “snake toxins inhibition” OR “snake toxins neutralization” OR “viper” OR “viperidae” OR “crotalinae” OR “viperinae” OR “elapidae” OR “pit-viper” OR “bothrops” OR “jararaca” OR “crotalus” OR “micrurus” OR “lachesis” OR “cobra” OR “naja” OR “bitis” OR “vipera” OR “daboia” OR “trimeresus”).All abstracts and/or full-text data were considered, without language restriction. Then, the publications covering ethnobotanical and/or pharmacological studies of antiophidic plants were selected and carefully analyzed. With the information gathered in these studies, the actual scenario of the use of plants against snake venom was pointed out. Main botanical families used, main countries where antiophidic plants are reported, and mode of use mostly employed in folk medicine were described. Regarding studies of pharmacological evidence, the snake species that were most studied, which plant species were tested and presented positive results, correlating with those species that also presented record of ethnopharmacological use, were also reported.The accepted botanical name of each medicinal plant listed was confirmed in at least 2 botanical databases among the following ones: Flora do Brasil (http://www.floradobrasil.jbrj.gov.br/), Tropicos (http://www.tropicos.org/), The Plant List (http://www.theplantlist.org/), and NCBI Taxonomy Browser (https://www.ncbi.nlm.nih.gov/taxonomy). In some cases, where the same species was considered as different ones (different synonyms used) in different papers, the accepted name according to the botanical databases mentioned above was used in the present review, bringing the synonym used in the original work between parenthesis.


## 3. Medicinal Plants as a Popular Source of Antidotes for Snakebites: Traditional Use

According to the literature search performed, a lot of ethnopharmacological studies showing medicinal plants claimed as antiophidic were found. A summary of these vegetal species can be observed in Table1.Table 1
List of medicinal plants used against snakebites.
Plant name
Countries
Parts used
Use
Reference(s)
Acanthaceae
Acanthus arboreus
Sri Lanka
ND
I
[18]
Andrographis echioides (syn. Indoneesiella echioides)
India
Shoot
ND
[19]
Andrographis lineata
India
Flower, leaf
I
[20]
Andrographis paniculata
India
Leaf, whole plant
I, E
[16, 20–26]
Barleria cristata
India, Pakistan
Leaf, root, seed, whole plant
E
[17, 19, 25]
Barleria lupulina
Sri Lanka
ND
I
[18]
Blechum pyramidatum
Nicaragua
Leaf, whole plant
I
[27]
Blepharis maderaspatensis
India
Leaf
I
[28]
Clinacanthus nutans
India
Leaf
E
[20]
Dicliptera paniculata(syn.Peristrophe paniculata)
India
Root, whole plant
I, E
[24, 25]
Fittonia albivenis
Peru
Aerial parts
E
[29]
Hygrophila auriculata
India, Sri Lanka
Seed
I
[18, 23]
Justicia adhatoda(syn.Adhatoda vasica)
India, Pakistan, Sri Lanka
Flower, leaf, root
I, E
[16–18, 30]
Justicia calyculata
Kenya
Aerial parts
E
[31]
Justicia gendarussa
Bangladesh
Leaf
I
[30, 32]
Justicia japonica(syn. Justicia simplex)
India
Leaf
I
[23]
Justicia
p
e
c
t
o
r
a
l
i
s
∗
Brazil
Leaf
I
[33]
Justicia procumbens
Sri Lanka
ND
E
[18]
Justicia
s
e
c
u
n
d
a
#
Colombia
Flower, leaf, root, whole plant
I, E
[34, 35]
Rhinacanthus nasutus
India
Leaf, root
I, E
[16, 28]
Thunbergia alata
Colombia
Flower, leaf
E
[34]
Trichanthera gigantea
Colombia
Leaf, root
E
[34]
Acoraceae
Acorus calamus
Bangladesh, India, Pakistan, Sri Lanka
Rhizome, root
I, E
[17, 18, 20, 22, 25, 32, 36]
Adoxaceae
Sambucus nigra
Spain
Flower
E
[37]
Amaranthaceae
Achyranthes aspera(syn.Achyranthes porphyristachya)#
Bangladesh, Colombia, India
Fruit, inflorescence, leaf, root, seed, stem, whole plant
I, E
[17, 20, 22, 23, 25, 28, 30, 32, 35, 38–41]
Aerva lanata
India, Sri Lanka
Rhizome
I
[18, 20]
Aerva sanguinolenta
Bangladesh
Leaf
ND
[30]
Alternanthera albotomentosa
Colombia
Leaf
E
[34]
Alternanthera brasiliana
Brazil
Flower
I
[33]
Alternanthera sessilis
Sri Lanka
ND
I
[18]
Amaranthus blitum
India
Root
I
[25]
Amaranthus dubius
Colombia
Fruit peel, leaf, root, seed
E
[34]
Amaranthus polygonoides
Sri Lanka
ND
E
[18]
Amaranthus spinosus
India
Leaf, root, stem, whole plant
E
[17, 19, 32, 42]
Amaranthus viridis
Bangladesh, India, Pakistan, Sri Lanka
Leaf, stem, whole plant
E
[17, 18, 30, 42, 43]
Chenopodium album
Bangladesh, India, Pakistan
Fruit, root, whole plant
E
[17, 32, 41]
Cyathula tomentosa
India
Leaf
ND
[19]
Dysphania ambrosioides(syn.Chenopodium ambrosioides)
Colombia
Whole plant
E
[34]
Amaryllidaceae
Allium ascalonicum
Sri Lanka
ND
I
[18]
Allium
c
e
p
a
∗
Bangladesh, Colombia, India, Kenya
Bulb, latex, leaf
E
[20, 25, 31, 32, 34, 40]
Allium
s
a
t
i
v
u
m
∗
Colombia, India, Sri Lanka, Spain
Bulb, inflorescence, leaf
I, E
[18, 22, 23, 37, 44]
Ammocharis tinneana
Kenya
Latex
ND
[31]
Crinum asiaticum
Sri Lanka
ND
E
[18]
Crinum latifolium
Sri Lanka
ND
E
[18]
Hymenocallis littoralis
Nicaragua
Leaf, root
I, E
[27]
Anacardiaceae
Anacardium
o
c
c
i
d
e
n
t
a
l
e
∗
India, Nicaragua
Bark, fruit, leaf, root
I, E
[27, 45]
Buchanania cochinchinensis(syn.Buchanania lanzan)
India
Bark
E
[24, 38]
Mangifera
i
n
d
i
c
a
∗
Bangladesh, Pakistan, Sri Lanka
Leaf
E
[17, 18, 32]
Mangifera minor
Papua New Guinea
Bark
I
[46]
Pistacia chinensis
Pakistan
Gall
E
[17]
Pistacia chinensissubsp.integerrima∗
India, Pakistan
Gall, leaf
E
[17, 19]
Semecarpus anacardium
India
Root
I
[20]
Semecarpus coriacea
Sri Lanka
ND
E
[18]
Spondias dulcis
Sri Lanka
ND
E
[18]
Spondias
m
o
m
b
i
n
∗
Peru
Bark
ND
[29]
Tapirira guianensis
Colombia
Oil
E
[34]
Annonaceae
Annona
m
o
n
t
a
n
a
#
Brazil
Leaf
I
[33, 47]
Annona muricata
Brazil
Leaf
ND
[48]
Annona
s
e
n
e
g
a
l
e
n
s
i
s
∗
Kenya
Leaf
I, E
[31]
Annona squamosa
Bangladesh, India
Bark, fruit
I, E
[23, 32]
Polyalthia longifolia
Bangladesh
Whole plant
ND
[30]
Uvaria scheffleri
Kenya
Leaf, root
E
[31]
Apiaceae
Centella asiatica
Sri Lanka
ND
E
[18]
Conium maculatum
Spain
Leaf
E
[37]
Coriandrum sativum
Sri Lanka
ND
I
[18]
Eryngium bourgatii
Spain
Aerial parts, root
E
[37]
Eryngium campestre
Spain
Aerial parts, root
E
[37]
Eryngium foetidum
Nicaragua, Sri Lanka
Leaf
I, E
[18, 27]
Steganotaenia araliacea
Kenya
Root
E
[31]
Trachyspermum ammi
Sri Lanka
ND
I
[18]
Trachyspermum roxburghianum
Sri Lanka
ND
I
[18]
Apocynaceae
Allamanda
c
a
t
h
a
r
t
i
c
a
∗
Colombia
Aerial parts, branch, leaf, stem
I, E
[35, 44]
Alstonia scholaris
Bangladesh, India, Sri Lanka
Bark, flower, latex, leaf, root
I, E
[18, 19, 32]
Alstonia venenata
Sri Lanka
ND
E
[18]
Asclepias curassavica
Nicaragua
Bark, flower, latex, leaf, root, whole plant
I, E
[27]
Blepharodon mucronatum
Nicaragua
Leaf, whole plant
I, E
[27]
Calotropis acia
Sri Lanka
ND
I, E
[18]
Calotropis gigantea
Bangladesh, India, Sri Lanka
Latex, leaf, root
I, E
[16, 18, 20, 22, 23, 28, 32, 38, 49]
Calotropis
p
r
o
c
e
r
a
∗
Bangladesh, India, Pakistan
Flower, latex, leaf, root, shoot
I, E
[17, 25, 32, 38, 40, 41]
Cascabela thevetia(syn.Thevetia peruviana)
Brazil
Bark, seed
E
[50]
Catharanthus roseus
Bangladesh, Colombia
Flower, leaf
I, E
[32, 34]
Cerbera floribunda
Papua New Guinea
Latex
E
[46]
Cerbera odollam
Sri Lanka
ND
E
[18]
Cryptolepis dubia(syn.Cryptolepis buchanani)
India, Sri Lanka
Root
ND
[18, 19]
Cynanchum viminale(syn.Sarcostemma viminale)
India
Whole plant
E
[38]
Dregea volubilis(syn.Wattakaka volubilis)
India, Sri Lanka
Root
I, E
[18, 23]
Echidnopsis dammanniana
Ethiopia
Stem
E
[51]
Echites umbellatus
Nicaragua
Root
I
[27]
Gymnema sylvestre
India
Leaf, root
I, E
[19, 20, 22, 23, 28, 52]
Hemidesmus
i
n
d
i
c
u
s
∗
Bangladesh, India, Sri Lanka
Root, whole plant
I, E
[18, 20, 22, 25, 28, 32, 38]
Holarrhena pubescens(syn.Holarrhena antidysenterica)
Bangladesh, India
Bark, root, stem
I, E
[24, 32, 38]
Hoya ovalifolia
Sri Lanka
ND
I
[18]
Hunteria zeylanica
Sri Lanka
ND
E
[18]
Ichnocarpus frutescens
Bangladesh
Root
I
[32]
Nerium oleander(syn.Nerium indicum)#
India, Pakistan, Sri Lanka
Leaf, root, seed
E
[17, 18, 20, 28, 41, 53]
Odontadenia puncticulosa
Nicaragua
Leaf
I
[27]
Pergularia daemia
India, Namibia
Leaf
I
[19, 28, 54]
Rauvolfia serpentina
Bangladesh, India, Sri Lanka
Flower, leaf, rhizome, root, seed
I, E
[18, 20, 22, 28, 30, 32]
Rauvolfia tetraphylla(syn.Rauvolfia canescens)#
Bangladesh, India
Root
E
[16, 30]
Tabernaemontana dichotoma
Sri Lanka
ND
E
[18]
Tabernaemontana divaricata
Sri Lanka
ND
I
[18]
Tabernaemontana sananho
Peru
Leaf
E
[29]
Tylophora
i
n
d
i
c
a
∗
Bangladesh, India
Leaf
I
[23, 30, 32]
Tylophora longifolia
India
Flower, leaf
ND
[20]
Vincetoxicum hirundinaria
India
Root
ND
[19]
Willughbeia edulis
Bangladesh
Stem
E
[32]
Wrightia antidysenterica
Sri Lanka
ND
E
[18]
Wrightia arborea
India
Bark
ND
[19]
Wrightia tinctoria
India
Leaf
ND
[38]
Aponogetonaceae
Aponogeton crispus
Sri Lanka
ND
E
[18]
Araceae
Alocasia
c
u
c
u
l
l
a
t
a
#
Colombia
Rhizome, root
E
[35, 44]
Amorphophallus commutatus
India
Tuber
ND
[55]
Amorphophallus paeoniifolius
Sri Lanka
ND
I
[18]
Anaphyllum beddomei
India
Rhizome
E
[16]
Anthurium marmoratum
Colombia
Branch, leaf, stem
E
[35]
Arisaema concinnum
India
Fruit, tuber
ND
[19]
Arisaema flavum
Pakistan
Rhizome
ND
[17]
Arisaema jacquemontii
India, Pakistan
Flower, leaf, tuber
ND
[17, 19, 56]
Arisaema tortuosum
India
Bulb, tuber
I
[38, 55]
Caladium bicolor
Peru
Tuber
E
[57]
Dieffenbachia
l
o
n
g
i
s
p
a
t
h
a
#
Colombia
Whole plant
I, E
[35]
Dieffenbachia parlatorei
Colombia
Root
E
[44]
Dracontium
c
r
o
a
t
i
i
∗
Colombia
Rhizome
I, E
[35]
Dracontium spruceanum
Colombia, Peru
Stem, tuber, root
E
[29, 34, 44, 57]
Dracunculus vulgaris
Spain
Bulb, flower
E
[37]
Homalomena aromatica
Bangladesh
Rhizome
E
[32]
Homalomena peltata
Colombia
Leaf
E
[44]
Homalomena picturata
Colombia
Leaf
E
[34]
Lasia spinosa
Sri Lanka
ND
E
[18]
Philodendron deltoideum
Peru
Aerial parts
I, E
[29]
Philodendron hederaceum
Nicaragua
Leaf, stem
I, E
[27]
Philodendron heleniae
Colombia
Leaf
E
[44]
Philodendron
m
e
g
a
l
o
p
h
y
l
l
u
m
∗
Brazil
Vine
I
[33]
Philodendron
t
r
i
p
a
r
t
i
t
u
m
∗
Colombia
Branch, leaf
E
[35]
Pothos scandens
Sri Lanka
ND
I
[18]
Rhodospatha oblongata
Colombia
Rhizome
E
[35]
Sauromatum venosum
India, Pakistan
Leaf, tuber
I, E
[17, 38]
Typhonium roxburghii
Sri Lanka
ND
I
[18]
Xanthosoma poeppigii
Peru
Stem
E
[57]
Araliaceae
Osmoxylon micranthum
Papua New Guinea
Latex
E
[46]
Arecaceae
Areca catechu
Sri Lanka
ND
E
[18]
Caryota urens
Sri Lanka
ND
I
[18]
Cocos nucifera
Sri Lanka
ND
I
[18]
Corypha umbraculifera
Sri Lanka
ND
E
[18]
Euterpe edulis
Brazil
Latex
E
[50]
Euterpe oleracea
Brazil
Fruit
E
[33]
Phoenix pusilla
Sri Lanka
ND
I
[18]
Syagrus coronata
Brazil
Bark
ND
[47]
Aristolochiaceae
Aristolochia birostris
Brazil
Whole plant
I
[47]
Aristolochia
b
r
a
c
t
e
o
l
a
t
a
∗
India, Sri Lanka
Fruit, leaf, whole plant
I, E
[18, 23, 38, 55]
Aristolochia clematitis
Serbia
Rhizome
ND
[58]
Aristolochia cordiflora
Colombia
Leaf, stem
I, E
[34, 44]
Aristolochia
g
r
a
n
d
i
f
l
o
r
a
#
Colombia
Whole plant
I, E
[35]
Aristolochia
i
n
d
i
c
a
∗
Bangladesh, India
Leaf, root, whole plant
I, E
[16, 20, 22, 23, 28, 30, 32]
Aristolochia ovalifolia
Mexico
Root
ND
[59]
Aristolochia
p
i
l
o
s
a
#
Colombia
Root
I, E
[35]
Aristolochia tagala
India
Whole plant
I, E
[16]
Aristolochia trilobata
Brazil, Nicaragua
Leaf, root, whole plant
I, E
[27, 50]
Thottea siliquosa
India
Leaf, root
E
[16, 26]
Asparagaceae
Asparagus racemosus
Bangladesh, Sri Lanka
Leaf, root
E
[18, 30, 32]
Drimia indica(syn.Urginea indica)
India
Bulb
E
[25]
Peliosanthes teta
Bangladesh
Root
E
[32]
Sansevieria parva
Kenya
Latex
E
[31]
Sansevieria roxburghiana
India
Rhizome
I
[23]
Sansevieria trifasciata
Bangladesh, Colombia
Aerial parts, whole plant
E
[30, 34, 60]
Sansevieria zeylanica
Sri Lanka
ND
E
[18]
Aspleniaceae
Asplenium dalhousiae(syn.Ceterach dalhousiae)
Pakistan
Leaf
ND
[17]
Asteraceae
Achillea millefolium
India
Whole plant
I
[20]
Acmella paniculata(syn. Spilanthes paniculata)
Sri Lanka
ND
I
[18]
Adenostemma fosbergii
Ecuador
Leaf
I
[61]
Adenostemma
l
a
v
e
n
i
a
#
Colombia
Whole plant
E
[35]
Ageratum
c
o
n
y
z
o
i
d
e
s
#
Colombia, India, Bangladesh
Flower, leaf, root
E
[19, 24, 32, 34]
Ageratum houstonianum
Pakistan
Inflorescence, leaf
E
[17]
Ambrosia peruviana(syn.Ambrosia cumanensis)
Colombia
Aerial parts, whole plant
I, E
[34, 44]
Artemisia maritima
Pakistan
Whole plant
E
[17]
Artemisia scoparia
India, Pakistan
Whole plant
E
[17, 40]
Austroeupatorium inulifolium
Colombia
Leaf
E
[34, 44]
Ayapana triplinervis(syn.Eupatorium ayapana,Eupatorium triplinerve)
Brazil
Leaf
I
[33, 50]
Baccharis inamoena(syn. Baccharis trinervis)
Colombia
Aerial parts, whole plant
E
[34]
Baccharoides anthelmintica(syn.Centratherum anthelminticum)
India
Seed
ND
[26]
Bidens biternata
India
Leaf
E
[25]
Bidens pilosa
Kenya
Leaf
E
[31]
Blumea axillaris
Sri Lanka
ND
I
[18]
Blumea brevipes(syn.Laggera brevipes)
Kenya
Root
ND
[31]
Calendula officinalis
India
Flower
I
[20, 28]
Chromolaena odorata
Colombia
Whole plant
E
[34]
Clibadium
s
y
l
v
e
s
t
r
e
#
Colombia
Whole plant
I, E
[35]
Conyza sumatrensis
Kenya
Leaf
I
[31]
Cyanthillium cinereum
Sri Lanka
ND
E
[18]
Eclipta prostrata(syn. Eclipta alba)∗
Bangladesh, India, Pakistan, Sri Lanka
Leaf, whole plant
I, E
[17, 18, 20, 28, 30, 40]
Elephantopus scaber
Sri Lanka
ND
E
[18]
Emilia sonchifolia
Bangladesh, Colombia, India, Sri Lanka
Leaf, whole plant
I, E
[16, 18, 30, 34]
Erechtites
v
a
l
e
r
i
a
n
i
f
o
l
i
a
#
Colombia
Branch, leaf, stem
I, E
[35]
Gnaphalium purpureum
Sri Lanka
ND
I, E
[18]
Gynura hispida
Sri Lanka
ND
I
[18]
Helianthus annuus
India
Seed
E
[20]
Inula helenium
Serbia
Root
E
[58]
Laggera alata
Sri Lanka
ND
E
[18]
Linzia glabra (syn. Vernonia glabra)
Kenya
Leaf
E
[31]
Microglossa pyrifolia
Kenya
Leaf
E
[31]
Mikania cordata
Bangladesh
Leaf
E
[32]
Mikania cordifolia
Nicaragua
Leaf, stem, whole plant
I, E
[27]
Mikania
g
u
a
c
o
#
Colombia, Nicaragua
Leaf, stem, whole plant
I, E
[27, 35, 44]
Neurolaena
l
o
b
a
t
a
∗
Colombia, Nicaragua
Aerial parts, branch, leaf, stem
I, E
[27, 35, 44]
Pentanema indicum
India, Sri Lanka
Leaf, root
I
[18, 23]
Pluchea
i
n
d
i
c
a
∗
India
Flower, seed
I, E
[20]
Pseudelephantopus
s
p
i
c
a
t
u
s
∗
Colombia
Whole plant
E
[44]
Saussurea simpsoniana
India
Flower
ND
[19]
Senecio chrysanthemoides
Pakistan
Whole plant
E
[17]
Seriphidium brevifolium(syn.Artemisia brevifolia)
Pakistan
Flower, leaf
E
[17]
Solanecio mannii
Kenya
Leaf
E
[31]
Sphaeranthus africanus
Sri Lanka
ND
I
[18]
Sphaeranthus indicus
Sri Lanka
ND
I
[18]
Sphagneticola trilobata
Nicaragua
Flower, leaf, stem, whole plant
I
[27]
Tagetes minuta
Kenya
Leaf
E
[31]
Taraxacum officinale
Colombia, Pakistan
Leaf, root, whole plant
I, E
[17, 34]
Tithonia diversifolia
Colombia, Kenya
Leaf, whole plant
I, E
[31, 34]
Tricholepis glaberrima
India
Root
ND
[19]
Verbesina gigantea
Colombia
Root, stem
I, E
[34]
Vernonanthura patens
Colombia
Whole plant
E
[34]
Vernonia zeylanicum
Sri Lanka
ND
I, E
[18]
Wedelia calendulacea
India
Leaf
I
[20]
Wollastonia biflora(syn.Wedelia biflora)
Sri Lanka
ND
E
[18]
Xanthium strumarium
Pakistan
Aerial parts
E
[17]
Balsaminaceae
Impatiens balsamina
Colombia
Flower
I, E
[34]
Begoniaceae
Begonia annulata(syn. Begonia barbata)
Bangladesh
Leaf, stem
E
[32]
Berberidaceae
Dysosma pleiantha
China, Taiwan
Rhizome
ND
[62]
Betulaceae
Betula alnoides
India
Bark, leaf
ND
[19]
Bignoniaceae
Crescentia
c
u
j
e
t
e
#
Colombia
Fruit
I
[35]
Dolichandra unguis-cati(syn.Macfadyena unguis-cati)#
Colombia
Whole plant
E
[35]
Handroanthus barbatus(syn. Tabebuia barbata)
Brazil
Leaf
I
[33]
Mansoa alliacea
Peru
Bark, root
I
[57]
Oroxylum indicum
Bangladesh, Sri Lanka
Bark
E
[18, 32]
Stereospermum chelonoides
Sri Lanka
ND
I
[18]
Stereospermum colais
Sri Lanka
ND
E
[18]
Tabebuia
r
o
s
e
a
∗
Colombia
Bark
I, E
[35]
Bixaceae
Bixa
o
r
e
l
l
a
n
a
∗
Bangladesh, Colombia, Nicaragua
Branch, fruit, latex, leaf, root, stem
I, E
[27, 32]
Cochlospermum vitifolium
Colombia
Aerial parts
E
[34]
Boraginaceae
Cordia dichotoma(syn.Cordia obliqua)
Pakistan
Bark, fruit
ND
[17]
Cordia spinescens(syn.Varronia spinescens)
Colombia
Leaf
E
[34]
Cynoglossum zeylanicum
India
Root
I
[63]
Echium vulgare
Spain
Aerial parts
ND
[37]
Ehretia microphylla(syn. Ehretia buxifolia)
India, Sri Lanka
Root
I, E
[18, 20]
Heliotropium europaeum
Pakistan
Whole plant
E
[17]
Heliotropium
i
n
d
i
c
u
m
#
Nicaragua
Leaf, whole plant
I
[27]
Tournefortia
c
u
s
p
i
d
a
t
a
#
Colombia
Branch, leaf, stem
E
[35]
Trichodesma
i
n
d
i
c
u
m
∗
Pakistan
Leaf, root
ND
[17]
Trichodesma zeylanicum
India
Root
I, E
[20]
Brassicaceae
Brassica juncea
Sri Lanka
ND
E
[18]
Brassica rapa (syn. Brassica campestris)
India
ND
E
[25]
Lepidium virginicum
Colombia
Whole plant
E
[34]
Bromeliaceae
Ananas comosus
Nicaragua, Sri Lanka
Flower, leaf, root
I, E
[18, 27]
Bromelia pinguin
Nicaragua
Leaf
I, E
[27]
Burseraceae
Boswellia serrata
India
Bark
I
[24]
Bursera simaruba
Nicaragua
Bark, whole plant
I
[27]
Canarium zeylanicum
Sri Lanka
ND
E
[18]
Cactaceae
Opuntia ficus-indica(syn.Opuntia vulgaris)
India
Root
ND
[25]
Pereskia
b
l
e
o
#
Colombia
Leaf, stem
E
[35]
Calophyllaceae
Calophyllum inophyllum
Sri Lanka
ND
E
[18]
Mesua ferrea
Sri Lanka
ND
I, E
[18]
Campanulaceae
Hippobroma longiflora
Nicaragua
Leaf, root, whole plant
I, E
[27]
Cannabaceae
Cannabis sativa
India, Sri Lanka
ND
I
[18, 40]
Cannaceae
Canna indica
Sri Lanka
ND
E
[18]
Capparaceae
Capparis decidua
Pakistan
Flower, shoot
E
[17]
Capparis moonii
Sri Lanka
ND
I
[18]
Capparis roxburghii
Sri Lanka
ND
E
[18]
Capparis zeylanica
Sri Lanka
ND
I, E
[18]
Carica
p
a
p
a
y
a
#
India
Fruit
ND
[41]
Crateva adansonii
Sri Lanka
ND
I
[18]
Crateva tapia(syn.Crateva benthamii)#
Brazil
Leaf
E
[33]
Cynophalla flexuosa(syn.Capparis flexuosa)
Brazil
Bark
I
[64]
Caprifoliaceae
Nardostachys jatamansi
India
Root
ND
[19]
Valeriana jatamansi
Pakistan, Sri Lanka
Root
I, E
[17, 18]
Celastraceae
Cassine glauca
India, Sri Lanka
Leaf
I
[18, 19]
Celastrus paniculatus
India
Bark, root, seed
I
[19, 38]
Gymnosporia emarginata
Sri Lanka
ND
I
[18]
Parnassia nubicola
India
Tuber
ND
[19]
Chrysobalanaceae
Parinari capensis
Namibia
Root
ND
[65]
Cleomaceae
Cleome gynandra
Sri Lanka
ND
E
[18]
Cleome viscosa
Sri Lanka
ND
I
[18]
Clusiaceae
Garcinia morella
Sri Lanka
ND
I, E
[18]
Garcinia xanthochymus
Sri Lanka
ND
I, E
[18]
Colchicaceae
Gloriosa
s
u
p
e
r
b
a
∗
India, Pakistan, Sri Lanka
Tuber
I, E
[17, 18, 20, 28, 38, 40]
Combretaceae
Anogeissus latifolia
Bangladesh, India
Bark, whole plant
I, E
[25, 30, 38]
Combretum collinum
Kenya
Root
E
[31]
Combretum
m
o
l
l
e
∗
Kenya
Bark, root
I
[31]
Getonia floribunda (syn. Calycopteris floribunda)
Bangladesh
Root
E
[32]
Terminalia
a
r
j
u
n
a
∗
Bangladesh, India
Bark
I, E
[20, 32]
Terminalia bellirica
Sri Lanka
ND
I
[18]
Terminalia chebula
Sri Lanka
ND
I
[18]
Commelinaceae
Callisia gracilis
Colombia
Flower, leaf
I, E
[34]
Commelina benghalensis
India, Sri Lanka
Root
ND
[18, 42]
Connaraceae
Connarus
f
a
v
o
s
u
s
∗
Brazil
Bark
I
[33]
Connarus monocarpus
Sri Lanka
ND
E
[18]
Convolvulaceae
Argyreia nervosa(syn.Argyreia speciosa)
India
Root, seed
ND
[19]
Argyreia populifolia
Sri Lanka
ND
I
[18]
Cuscuta reflexa
Sri Lanka
ND
E
[18]
Dichondra repens
Kenya
Leaf
E
[31]
Evolvulus alsinoides
India, Sri Lanka
Root
I
[18, 23]
Ipomoea alba
Sri Lanka
ND
E
[18]
Ipomoea aquatica
Bangladesh
Leaf, whole plant
ND
[30]
Ipomoea asarifolia
Sri Lanka
ND
I, E
[18]
Ipomoea
c
a
i
r
i
c
a
#
Colombia
Branch, leaf, stem
E
[35]
Ipomoea mauritiana
Nicaragua
Leaf
I, E
[27]
Ipomoea pes-caprae
Nicaragua
Leaf, seed
I
[27]
Ipomoea pes-tigridis
India, Sri Lanka
Root
I, E
[18, 19, 24, 39]
Ipomoea setifera
Nicaragua
Leaf
I, E
[27]
Ipomoea triloba
Sri Lanka
ND
I
[18]
Operculina pteripes
Nicaragua
Leaf
E
[27]
Rivea hypocrateriformis
India
ND
I
[24]
Cornaceae
Alangium salviifolium
India
Bark
I
[20, 23]
Costaceae
Cheilocostus speciosus (syn. Costus speciosus)
Bangladesh, India, Sri Lanka
Bulb, leaf, stem, root, tuber
I, E
[18, 19, 32, 55]
Costus
g
u
a
n
a
i
e
n
s
i
s
#
Colombia
Stem
I, E
[35]
Costus
l
a
s
i
u
s
∗
Colombia
Branch, leaf, stem
I, E
[35]
Costus lima
Colombia
Stem
E
[34]
Crassulaceae
Bryophyllum pinnatum(syn.Kalanchoe pinnata)∗
India
Leaf
ND
[22, 42]
Kalanchoe laciniata (syn. Kalanchoe brasiliensis)∗
Brazil
Leaf
E
[33]
Cucurbitaceae
Benincasa hispida
Sri Lanka
ND
E
[18]
Citrullus
c
o
l
o
c
y
n
t
h
i
s
∗
India, Pakistan
Fruit, root
ND
[17, 40, 41]
Coccinia grandis
Pakistan, Sri Lanka
Root
I, E
[17, 18]
Corallocarpus epigaeus
India
Tuber
I
[38]
Cucumis melo
Sri Lanka
ND
I
[18]
Cucurbita pepo
Spain
Flower
E
[37]
Diplocyclos palmatus
India, Sri Lanka
Leaf, tuber
I, E
[18, 23, 66]
Fevillea cordifolia
Colombia, Nicaragua
Seed, whole plant
I, E
[27, 35]
Lagenaria
s
i
c
e
r
a
r
i
a
#
Sri Lanka
ND
E
[18]
Luffa acutangula
India, Sri Lanka
Fruit, whole plant
I, E
[18, 19, 38]
Momordica balsamina
India
ND
ND
[40]
Momordica
c
h
a
r
a
n
t
i
a
∗
Colombia, India, Nicaragua, Sri Lanka
Aerial parts, branch, flower, fruit, leaf, stem, whole plant
I, E
[18, 20, 27, 34, 35]
Momordica dioica
Sri Lanka
ND
E
[18]
Sicydium tamnifolium
Mexico
Root
ND
[59]
Trichosanthes cucumerina
India, Sri Lanka
Leaf
I
[18, 38]
Trichosanthes tricuspidata
Bangladesh
Root
I
Cycadaceae
Cycas pectinata
Bangladesh
Flower
E
[32]
Cycas revoluta
Bangladesh
Whole plant
ND
[30]
Cyclanthaceae
Cyclanthus bipartitus
Peru
Heart
E
[57]
Cyperaceae
Cyperus kyllingia
Sri Lanka
ND
I
[18]
Cyperus rotundus
Bangladesh, India, Pakistan, Sri Lanka
Bulb, flower, leaf, rhizome, root, tuber
I, E
[17, 18, 20, 28, 32, 39]
Kyllinga odorata (syn. Kyllinga monocephala)
India
ND
ND
[40]
Dilleniaceae
Tetracera sarmentosa
Sri Lanka
ND
I, E
[18]
Dioscoreaceae
Dioscorea oppositifolia
Sri Lanka
ND
I
[18]
Dioscorea pentaphylla
India
Tuber
I
[38, 55]
Dipterocarpaceae
Dipterocarpus lowii
Sri Lanka
ND
I
[18]
Dipterocarpus zeylanicus
Sri Lanka
ND
E
[18]
Droseraceae
Drosera burmannii
Sri Lanka
ND
I, E
[18]
Drosera indica
Sri Lanka
ND
E
[18]
Ebenaceae
Diospyros kaki
Malaysia
Fruit
I
[67]
Diospyros melanoxylon
India
Seed
E
[25]
Diospyros montana
India
Root
I
[38]
Diospyros vera(syn. Maba buxifolia)
Sri Lanka
ND
I, E
[18]
Euclea racemosa
Ethiopia
Leaf
I
[51]
Elaeagnaceae
Elaeagnus latifolia
Sri Lanka
ND
I, E
[18]
Ericaceae
Gaultheria trichophylla
India
Leaf
I
[66]
Erythroxylaceae
Erythroxylum monogynum
Sri Lanka
ND
E
[18]
Euphorbiaceae
Acalypha aristata (syn. Acalypha arvensis)
Nicaragua
Leaf, whole plant
I, E
[27]
Acalypha fimbriata
ND
ND
ND
[68]
Acalypha
i
n
d
i
c
a
∗
Bangladesh, India, Sri Lanka
Leaf, whole plant
E
[18, 20, 32]
Acalypha phleoides
Mexico
ND
ND
[68]
Acalypha wilkesiana(syn.Acalypha godseffiana)
Sri Lanka
ND
E
[18]
Agrostistachys hookeri
Sri Lanka
ND
E
[18]
Baliospermum solanifolium(syn.Baliospermum montanum)
India
Leaf, root, seed
E
[19, 32]
Cnidoscolus aconitifolius
Colombia
Leaf, whole plant
I, E
[34]
Croton tiglium
Sri Lanka
ND
E
[18]
Croton trinitatis
Colombia
Whole plant
E
[34]
Euphorbia antiquorum
Sri Lanka
ND
E
[18]
Euphorbia
h
i
r
t
a
∗
Bangladesh, Brazil, India
Latex, root, whole plant
I
[19, 20, 32, 47]
Euphorbia milii
Bangladesh
Whole plant
ND
[30]
Euphorbia neriifolia (syn. Euphorbia ligularia)
India, Sri Lanka
Latex, leaf, stem
I, E
[18, 19, 22, 38]
Euphorbia thymifolia
Nicaragua
Latex, leaf, whole plant
I
[27]
Euphorbia tirucalli
Sri Lanka
ND
I
[18]
Euphorbia tithymaloides(syn.Pedilanthus tithymaloides)
Sri Lanka
ND
I, E
[18]
Euphorbia tortilis
Sri Lanka
ND
E
[18]
Hura crepitans
Peru
Latex
E
[57]
Jatropha
c
u
r
c
a
s
∗
Brazil, Nepal
Latex, root, stem
I
[47, 64, 69, 70]
Jatropha
g
o
s
s
y
p
i
i
f
o
l
i
a
∗
Bangladesh, Brazil
Latex, leaf, stem
I, E
[32, 50]
Jatropha
m
o
l
l
i
s
s
i
m
a
∗
Brazil
Latex
ND
[47, 64]
Jatropha multifida
Sri Lanka
ND
E
[18]
Jatropha podagrica
Sri Lanka
ND
E
[18]
Jatropha ribifolia
Brazil
Latex
ND
[47]
Mallotus repandus
Sri Lanka
ND
E
[18]
Manihot esculenta
Brazil, Colombia, Nicaragua
Branch, leaf, root
I, E
[27, 33, 34]
Melanolepis multiglandulosa
Papua New Guinea
Latex
I
[46]
Phyllanthus
a
c
u
m
i
n
a
t
u
s
#
Colombia
Branch, leaf
I, E
[35]
Ricinus communis
Brazil, Pakistan, Sri Lanka
Fruit, latex, leaf, root, seed
I, E
[17, 18, 69, 71, 72]
Spirostachys africana
Namibia
Stem
ND
[65]
Tragia involucrata
India
Whole plant
I
[20, 28]
Trewia nudiflora
Bangladesh
Leaf
E
[32]
Fabaceae
Abrus
p
r
e
c
a
t
o
r
i
u
s
∗
Bangladesh, India
Leaf, root, stem
I, E
[20, 21, 28, 32, 38]
Abrus pulchellus
Sri Lanka
ND
E
[18]
Acacia caesia
Sri Lanka
ND
I, E
[18]
Acacia cornigera
Mexico
Root
ND
[59]
Acacia leucophloea
India
Bark
I, E
[20, 63]
Acacia mellifera
Namibia
ND
ND
[54]
Acacia nilotica
India
Leaf
I, E
[38]
Acacia torta
India
Bark
I
[63]
Acosmium panamense
Mexico
Bark
ND
[59]
Adenanthera pavonina
Sri Lanka
ND
I, E
[18]
Albizia
l
e
b
b
e
c
k
∗
Bangladesh, India, Pakistan, Sri Lanka
Bark, flower, fruit, leaf, seed
I, E
[16–18, 23, 32, 40]
Albizia procera
Bangladesh, Pakistan
Juicy parts, leaf, root
E
[17, 32]
Alysicarpus vaginalis
Sri Lanka
ND
I
[18]
Amburana cearensis
Brazil
Seed
ND
[71]
Bauhinia divaricata(syn.Bauhinia retusa)
India
Bark, flower, leaf
ND
[19]
Bauhinia guianensis
Nicaragua
Bark, stem
I, E
[27]
Bauhinia purpurea
India
Bark, flower, leaf
ND
[19]
Bauhinia racemosa
Sri Lanka
ND
E
[18]
Bauhinia
v
a
r
i
e
g
a
t
a
∗
Bangladesh, Sri Lanka
Bulb, stem
E
[18, 32]
Brownea
rosa-de-monte
∗
Colombia
Bark
I, E
[35]
Butea
m
o
n
o
s
p
e
r
m
a
∗
India
Bark, leaf, resin, seed
I, E
[24, 25, 38, 40, 41]
Caesalpinia bonduc
India, Nicaragua, Sri Lanka
Root, seed
I, E
[18, 20, 27, 38]
Caesalpinia coriaria
Sri Lanka
ND
E
[18]
Cajanus cajan
Bangladesh
Stem
E
[30, 32]
Canavalia gladiata
Sri Lanka
ND
E
[18]
Cassia
f
i
s
t
u
l
a
∗
Bangladesh, Brazil, India, Sri Lanka
Bark, fruit, leaf, root, seed
I, E
[18, 19, 24, 25, 32, 33, 38, 40]
Centrosema pubescens
Colombia
Whole plant
E
[34]
Clitoria ternatea
Bangladesh, India, Sri Lanka
Flower, leaf, root, seed
I, E
[16, 18, 19, 32, 38, 39, 42, 60]
Crotalaria laburnifolia
Sri Lanka
ND
E
[18]
Crotalaria verrucosa
India
Seed
I
[23]
Dalbergia melanoxylon
India
Bark
I
[20]
Deguelia amazonica (syn. Derris amazonica)
Brazil
Root
ND
[50]
Derris floribunda
Brazil
Root
ND
[50]
Desmodium
a
d
s
c
e
n
d
e
n
s
#
Colombia, Nicaragua
Leaf, root, whole plant
I, E
[27, 35]
Desmodium gangeticum
Bangladesh, India, Pakistan
Root, whole plant
I, E
[17, 32, 55]
Desmodium triflorum
Bangladesh, Sri Lanka
Shoot
I, E
[18, 32]
Dipteryx
o
d
o
r
a
t
a
#
Brazil
Seed
I
[33, 50]
Entada leptostachya
Kenya
Latex
E
[31]
Entada rheedii(syn. Entada pursaetha)
Bangladesh, India, Sri Lanka
Leaf, seed
I, E
[18, 32, 49]
Erythrina americana
Mexico
Leaf, seed
ND
[59]
Erythrina excelsa
India, Kenya
Bark, latex
ND
[20, 31]
Erythrina fusca
Sri Lanka
ND
I, E
[18]
Erythrina subumbrans
Sri Lanka
ND
I
[18]
Erythrina variegata
India
Bark
ND
[19]
Gliricidia sepium
Colombia
Leaf, stem
I, E
[34]
Glycine max
India
Seed
I
[20]
Glycyrrhiza glabra
Sri Lanka
ND
E
[18]
Humboldtia decurrens
India
Root
E
[16]
Humboldtia laurifolia
Sri Lanka
ND
E
[18]
Indigofera circinella
Kenya
Leaf
E
[31]
Indigofera suffruticosa
Colombia, Nicaragua
Aerial parts, seed, whole plant
I, E
[27, 34]
Indigofera
t
i
n
c
t
o
r
i
a
#
India
Root
I
[16]
Leucaena leucocephala
Sri Lanka
ND
E
[18]
Libidibia
f
e
r
r
e
a
#
Brazil
Seed
I
[33]
Machaerium ferox
Brazil
Leaf
E
[33]
Macrotyloma uniflorum
Sri Lanka
ND
I
[18]
Mimosa
p
u
d
i
c
a
∗
Bangladesh, India
Leaf, root, whole plant
I, E
[16, 19, 20, 22, 23, 28, 32]
Mucuna
p
r
u
r
i
e
n
s
#
Bangladesh, India, Nepal, Sri Lanka
Fruit, seed, stem, whole plant
I, E
[18, 19, 28, 32, 69]
Mucuna sloanei
Ecuador
Seed
I
[61]
Mucuna urens
Nicaragua
Seed
E
[27]
Parkinsonia aculeata
Brazil
Seed
ND
[47]
Pentaclethra
m
a
c
r
o
l
o
b
a
∗
Nicaragua
Bark
I, E
[27]
Plathymenia
r
e
t
i
c
u
l
a
t
a
∗
Brazil
Bark
I
[33]
Pongamia pinnata
Sri Lanka
ND
I, E
[18]
Pterocarpus santalinus
Sri Lanka
ND
E
[18]
Saraca asoca
Sri Lanka
ND
I
[18]
Senna alata(syn.Cassia alata)
India, Nicaragua, Sri Lanka
Flower, leaf, whole plant
I, E
[18, 20, 27, 28]
Senna
a
u
r
i
c
u
l
a
t
a
∗
Sri Lanka
ND
E
[18]
Senna
d
a
r
i
e
n
s
i
s
∗
Colombia
Whole plant
I, E
[35]
Senna hirsuta
Bangladesh
Leaf
E
[32]
Senna occidentalis (syn. Cassia occidentalis)
Bangladesh, India, Nicaragua, Sri Lanka
Leaf, root, whole plant
I, E
[18, 27, 32, 40]
Senna reticulata(syn.Cassia reticulata)
Brazil, Nicaragua
Leaf, root, whole plant
I
[27, 50]
Senna siamea
Kenya
Root
ND
[31]
Senna sophera(syn.Cassia sophera)
Bangladesh
Leaf, root
I
[30, 32]
Senna tora(syn. Cassia tora)
Bangladesh, India
Leaf, root, seed, stem
I, E
[20, 24, 25, 28, 32, 42]
Sesbania grandiflora
Sri Lanka
ND
I, E
[18]
Tadehagi triquetrum(syn.Desmodium triquetrum)
India
Whole plant
ND
[19]
Tamarindus
i
n
d
i
c
a
∗
Bangladesh, India, Sri Lanka
Seed, whole plant
I, E
[18, 22, 25, 32, 38]
Tephrosia purpurea
Bangladesh, India
Root, whole plant
I, E
[19, 20, 24, 32]
Trigonella foenum-graecum
Sri Lanka
ND
I
[18]
Uraria lagopodioides
India
Bark
I, E
[49]
Uraria picta
Bangladesh, India
Root, whole plant
I
[24, 30]
Vigna luteola
Colombia
Whole plant
E
[34]
Vigna radiata
Sri Lanka
ND
I
[18]
Gentianaceae
Chelonanthus alatus(syn.Irlbachia alata)#
Colombia
Branch, leaf
E
[35]
Enicostema
a
x
i
l
l
a
r
e
∗
India
Whole plant
I
[23, 45]
Fagraea ceilanica
Sri Lanka
ND
E
[18]
Hoppea dichotoma
India
Shoot
ND
[19]
Huperzia phlegmaria
Sri Lanka
ND
E
[18]
Potalia amara
Peru
Aerial parts
ND
[29]
Gesneriaceae
Columnea
p
u
l
c
h
e
r
r
i
m
a
#
Colombia
Whole plant
I, E
[35]
Columnea sanguinea(syn.Besleria sanguinea)#
Colombia
Whole plant
I, E
[35]
Episcia
d
i
a
n
t
h
i
f
l
o
r
a
#
Colombia
Whole plant
I, E
[35]
Gleicheniaceae
Gleichenella pectinata
Colombia
Whole plant
I
[34]
Haemodoraceae
Xiphidium
c
a
e
r
u
l
e
u
m
#
Colombia, Nicaragua, Peru
Leaf, stem, whole plant
I, E
[27, 35, 44, 57]
Heliconiaceae
Heliconia
c
u
r
t
i
s
p
a
t
h
a
∗
Colombia
Rhizome
E
[35]
Hydroleaceae
Hydrolea zeylanica
Sri Lanka
ND
I
[18]
Hymenophyllaceae
Trichomanes
e
l
e
g
a
n
s
∗
Colombia
Whole plant
E
[35]
Hypoxidaceae
Curculigo orchioides
Bangladesh, India
Bulb, leaf, rhizome
I
[32, 73]
Iridaceae
Iris kemaonensis
India
Rhizome
ND
[66]
Sisyrinchium micranthum
Colombia
Whole plant
E
[34]
Lamiaceae
Aegiphila
p
a
n
a
m
e
n
s
i
s
#
Colombia
Leaf, branch, stem
E
[35]
Anisochilus velutinus
Sri Lanka
ND
E
[18]
Anisomeles indica
India, Sri Lanka
Whole plant
ND
[18, 19]
Anisomeles malabarica
Bangladesh, India
Whole plant
I
[28, 30, 60]
Callicarpa tomentosa
Sri Lanka
ND
E
[18]
Clerodendrum cordatum(syn.Clerodendrum viscosum)
Bangladesh
Flower, leaf
E
[32]
Clerodendrum phlomidis
Sri Lanka
ND
E
[18]
Fuerstia africana
Kenya
Leaf
I
[31]
Gmelina arborea
Bangladesh
Root
I
[32]
Gmelina asiatica
Sri Lanka
ND
I, E
[18]
Hyptis
c
a
p
i
t
a
t
a
#
Colombia
Branch, leaf, stem
I, E
[35]
Hyptis suaveolens
Bangladesh
Leaf
E
[32]
Leonotis leonurus
South Africa
Flower, leaf
I
[74]
Leucas
a
s
p
e
r
a
∗
Bangladesh, India
Leaf, root, stem
I
[23, 24, 28, 30, 32]
Leucas
c
e
p
h
a
l
o
t
e
s
∗
India
Bark, leaf, whole plant
I, E
[19, 20, 40, 49]
Marsypianthes
c
h
a
m
a
e
d
r
y
s
∗
Brazil
Leaf
I
[33]
Mentha × piperita
Colombia
Leaf
E
[34]
Mentha pulegium
Colombia
Leaf
E
[34]
Ocimum
b
a
s
i
l
i
c
u
m
#
Bangladesh, Colombia, India
Branch, leaf, stem, whole plant
I, E
[20, 32, 35]
Ocimum campechianum(syn.Ocimum micranthum)
Colombia, Nicaragua
Aerial parts, leaf, whole plant
I, E
[27, 44]
Ocimum tenuiflorum(syn.Ocimum sanctum)∗
India, Sri Lanka
Leaf, root, whole plant
I, E
[16, 18, 20, 28, 40, 41]
Origanum vulgare
Serbia
Flower, leaf
ND
[58]
Plectranthus amboinicus
Sri Lanka
ND
I
[18]
Plectranthus hadiensis
Sri Lanka
ND
I
[18]
Plectranthus monostachyus
Brazil
Leaf
I
[33]
Pogostemon cablin
Malaysia
ND
ND
[75]
Pogostemon heyneanus
Sri Lanka
ND
E
[18]
Premna esculenta
Bangladesh
Leaf
E
[32]
Premna serratifolia(syn.Premna integrifolia)
Bangladesh
Leaf, root
I, E
[36]
Rosmarinus officinalis
Colombia
Whole plant
E
[34]
Rotheca serrata(syn.Clerodendrum serratum)
India
Leaf, root
ND
[19, 39]
Tectona grandis
India
Bark
I
[25]
Teucrium chamaedrys
Serbia
Flower
ND
[58]
Thymus vulgaris
India, Spain
Aerial parts, whole plant
I, E
[20, 37]
Vitex
n
e
g
u
n
d
o
∗
Bangladesh, India, Sri Lanka
Leaf, rhizome, root
I, E
[18, 20, 22, 32]
Vitex trifolia
India
Leaf
I
[28]
Volkameria eriophylla(syn. Clerodendrum eriophyllum)
Kenya
Leaf, root
ND
[76]
Lauraceae
Aniba parviflora(syn.Aniba fragrans)∗
Brazil
Bark
I
[33]
Cinnamomum verum
Sri Lanka
ND
I, E
[18]
Litsea glutinosa
Sri Lanka
ND
E
[18]
Litsea longifolia
Sri Lanka
ND
I, E
[18]
Persea macrantha
Sri Lanka
ND
E
[18]
Lecythidaceae
Careya arborea
Sri Lanka
ND
E
[18]
Couroupita guianensis
Bangladesh
Bark, leaf
ND
[30]
Linderniaceae
Lindernia
d
i
f
f
u
s
a
#
Colombia
Whole plant
E
[35]
Loganiaceae
Strychnos
nux-vomica
∗
India
Bark, root, seed
I, E
[16, 20, 49]
Strychnos potatorum
Sri Lanka
ND
E
[18]
Strychnos
x
i
n
g
u
e
n
s
i
s
∗
Colombia
Stem
E
[35]
Loranthaceae
Struthanthus cassythoides
Nicaragua
Leaf, whole plant
I, E
[27]
Struthanthus
o
r
b
i
c
u
l
a
r
i
s
∗
Colombia
Branch, leaf
E
[35]
Lycopodiaceae
Huperzia pulcherrima
Sri Lanka
ND
E
[18]
Lygodiaceae
Lygodium heterodoxum
Nicaragua
Leaf
I, E
[27]
Lygodium venustum
Colombia, Mexico, Nicaragua
Aerial parts, leaf, stem, whole plant
I, E
[27, 34, 59]
Lythraceae
Lawsonia inermis
India
Bark
ND
[25]
Punica granatum
India, Sri Lanka
Whole plant
I, E
[18, 20, 28]
Trapa natans(syn.Trapa bispinosa)
Sri Lanka
ND
I
[18]
Magnoliaceae
Magnolia champaca(syn.Michelia champaca)
Sri Lanka
ND
E
[18]
Malpighiaceae
Bronwenia cornifolia(syn.Banisteriopsis cornifolia)
Nicaragua
Bark, leaf, stem
E
[27]
Byrsonima crassifolia
Brazil, Nicaragua
Bark, leaf
I
[27, 47]
Stigmaphyllon puberum
Nicaragua
Leaf, stem
I, E
[27]
Malvaceae
Abelmoschus moschatus
Bangladesh, India, Sri Lanka
Fruit, leaf, seed
I, E
[18, 32, 38]
Abroma augusta
Bangladesh
Leaf, root, stem
E
[32]
Abutilon hirtum(syn. Abutilon heterotrichum)
Sri Lanka
ND
I, E
[18]
Abutilon indicum
India, Sri Lanka
Fruit, leaf
I
[18, 20]
Ceiba pentandra
Sri Lanka
ND
I
[18]
Corchorus trilocularis
Kenya
Leaf
E
[31]
Firmiana simplex(syn.Sterculia urens)
India
Bark, latex
I
[38, 55]
Gossypium arboreum
Sri Lanka
ND
E
[18]
Gossypium herbaceum
India
Seed
ND
[41]
Gossypium hirsutum
Brazil
Leaf
I
[33]
Grewia damine
Sri Lanka
ND
E
[18]
Grewia nervosa(syn.Microcos paniculata)
Sri Lanka
ND
E
[18]
Helicteres isora
Bangladesh, India
Fruit, root
I
[23, 25, 32]
Hibiscus rostellatus(syn.Hibiscus furcatus)
Sri Lanka
ND
E
[18]
Hibiscus surattensis
Sri Lanka
ND
E
[18]
Hibiscus tiliaceus
Mexico
Seed
ND
[59]
Melochia corchorifolia
Bangladesh, Sri Lanka
Leaf, whole plant
I, E
[18, 32]
Sida
a
c
u
t
a
#
Bangladesh, Colombia, India, Sri Lanka
Leaf, whole plant
I, E
[18, 32, 35, 39, 44]
Sida cordata
Sri Lanka
ND
I
[18]
Sida cordifolia
Bangladesh
Leaf
I
[32]
Sida rhombifolia
Bangladesh, Nicaragua, Sri Lanka
Leaf, stem
I, E
[18, 27, 32]
Thespesia populnea
Sri Lanka
ND
I
[18]
Triumfetta rhomboidea
Kenya
Root
E
[31]
Urena lobata
Bangladesh
Root
I
[32]
Wissadula periplocifolia
Bangladesh, Sri Lanka
Leaf, root
E
[18, 30, 60]
Marantaceae
Ischnosiphon rotundifolius
Brazil
Leaf
ND
[47]
Martyniaceae
Martynia annua
India, Sri Lanka
Fruit
E
[18, 25]
Melastomataceae
Osbeckia octandra
Sri Lanka
ND
E
[18]
Bellucia
d
i
c
h
o
t
o
m
a
∗
Brazil
Bark
I
[33]
Melastoma malabathricum
Bangladesh
Leaf
E
[32]
Memecylon umbellatum
India
Leaf
I
[63]
Meliaceae
Azadirachta indica
India, Sri Lanka
Bark, flower, latex, leaf, seed
I, E
[18, 20, 22, 28, 39–41]
Cipadessa baccifera
India
Leaf, root
I
[63]
Melia azedarach
India, Sri Lanka
Bark, leaf
I, E
[18, 41]
Munronia pinnata
Sri Lanka
ND
I, E
[18]
Menispermaceae
Cissampelos fasciculata
Colombia
Leaf
I
[44]
Cissampelos
p
a
r
e
i
r
a
∗
Bangladesh, India, Mexico, Nicaragua, Sri Lanka
Leaf, root, whole plant
I, E
[18, 19, 23, 25, 27, 32, 38, 55, 59]
Cocculus acuminatus
India
Stem
E
[16]
Cocculus hirsutus(syn.Cocculus villosus)
India
Leaf
I
[38, 40]
Coscinium fenestratum
Sri Lanka
ND
I
[18]
Cyclea peltata
Sri Lanka
ND
I
[18]
Odontocarya
t
e
n
a
c
i
s
s
i
m
a
#
Colombia
Whole plant
I, E
[35]
Tinospora cordifolia
Bangladesh, India, Sri Lanka
Fruit, root, stem
I
[18, 22, 23, 32]
Menyanthaceae
Nymphoides indica
Nicaragua, Sri Lanka
Leaf, root
I, E
[18, 27]
Monimiaceae
Hortonia angustifolia
Sri Lanka
ND
E
[18]
Moraceae
Artocarpus heterophyllus
Sri Lanka
ND
E
[18]
Artocarpus nobilis
Sri Lanka
ND
I, E
[18]
Broussonetia zeylanica
Sri Lanka
ND
I, E
[18]
Castilla
e
l
a
s
t
i
c
a
∗
Colombia
Branch, leaf, stem
I, E
[35]
Dorstenia contrajerva
Mexico, Nicaragua
Leaf, whole plant
I, E
[27, 59]
Ficus benghalensis
India
ND
ND
[40]
Ficus drupacea
Sri Lanka
ND
E
[18]
Ficus hispida
Sri Lanka
ND
E
[18]
Ficus
n
y
m
p
h
a
e
i
f
o
l
i
a
∗
Colombia
Branch, leaf, stem
I, E
[35]
Ficus racemosa
Bangladesh, India, Sri Lanka
Bark, shoot
I, E
[18, 32, 38]
Ficus religiosa
India, Sri Lanka
Bark
I, E
[18, 49]
Morus
a
l
b
a
∗
India
Leaf
I
[20]
Plecospermum spinosum
Sri Lanka
ND
I, E
[18]
Streblus asper
Bangladesh
Root
E
[32]
Moringaceae
Moringa
o
l
e
i
f
e
r
a
#
India, Sri Lanka
Bark, root, seed
I, E
[16, 18, 20, 22, 24, 28]
Musaceae
Ensete ventricosum(syn.Ensete edule)
Kenya
Latex
E
[31]
Musa × paradisíac
a
∗
Ecuador, India, Nicaragua, Sri Lanka
Bark, flower, latex
I, E
[18, 20, 27, 28, 61]
Myristicaceae
Myristica fragrans
Sri Lanka
ND
I
[18]
Myrtaceae
Myrcia bracteata(syn. Eugenia bracteata)
Sri Lanka
ND
I, E
[18]
Syzygium aromaticum
Sri Lanka
ND
I
[18]
Syzygium caryophyllatum
Sri Lanka
ND
E
[18]
Syzygium cumini(syn.Eugenia jambolana)
India, Pakistan, Sri Lanka
Bark, leaf
I
[17, 18, 20]
Syzygium zeylanicum
Sri Lanka
ND
E
[18]
Nelumbonaceae
Nelumbo nucifera
Sri Lanka
ND
I
[18]
Nepenthaceae
Nepenthes distillatoria
Sri Lanka
ND
E
[18]
Nyctaginaceae
Boerhavia coccinea
Pakistan
Whole plant
E
[17]
Boerhavia diffusa
Brazil, India, Sri Lanka
Leaf, root, whole plant
E
[18, 24, 25, 39, 41, 50]
Boerhavia procumbens
Pakistan
Leaf
E
[17]
Mirabilis jalapa
Bangladesh, Sri Lanka
Leaf
I, E
[18, 32]
Nymphaeaceae
Nymphaea nouchali
Sri Lanka
ND
E
[18]
Nymphaea pubescens
Sri Lanka
ND
I
[18]
Ochnaceae
Ochna jabotapita
Sri Lanka
ND
I
[18]
Sauvagesia erecta
Nicaragua
Whole plant
I, E
[27]
Oleaceae
Jasminum officinale
Sri Lanka
ND
E
[18]
Jasminum sambac
Sri Lanka
ND
E
[18]
Nyctanthes arbor-tristis
India, Sri Lanka
Root
I
[18, 49]
Olea europaea
Spain
Oil
ND
[37]
Opiliaceae
Opilia amentacea
Kenya
Root
E
[31]
Orchidaceae
Vanda tessellata
India
Root
E
[25]
Zeuxine regia
Sri Lanka
ND
E
[18]
Oxalidaceae
Averrhoa carambola
Sri Lanka
ND
I
[18]
Biophytum reinwardtii
Sri Lanka
ND
I
[18]
Oxalis corniculata
Bangladesh, Sri Lanka
Leaf
I, E
[18, 32]
Pandanaceae
Pandanus kaida
Sri Lanka
ND
I
[18]
Pandanus odorifer(syn.Pandanus odoratissimus)
India
Root
ND
[19]
Papaveraceae
Argemone mexicana
Bangladesh, India
Leaf, root, seed, stem
I, E
[20, 32, 38, 42]
Papilionaceae
Desmodium elegans
Pakistan
Root
E
[17, 53]
Passifloraceae
Adenia hondala
Sri Lanka
ND
E
[18]
Passiflora
q
u
a
d
r
a
n
g
u
l
a
r
i
s
∗
Colombia
Branch, leaf
E
[34, 35]
Phyllanthaceae
Antidesma bunius
India
Leaf
ND
[77]
Bridelia retusa
Sri Lanka
ND
I, E
[18]
Cleistanthus collinus
Sri Lanka
ND
I
[18]
Glochidion zeylanicum
Sri Lanka
ND
I
[18]
Margaritaria indica
Sri Lanka
ND
I, E
[18]
Phyllanthus acidus
India
Root
ND
[77]
Phyllanthus debilis
Sri Lanka
ND
I
[18]
Phyllanthus emblica(syn.Emblica officinalis)∗
Bangladesh, India, Sri Lanka
Bark, fruit, root
I, E
[18, 20, 22, 30]
Phyllanthus niruri
India
Flower
E
[20]
Phyllanthus reticulatus
India
Leaf
I
[20]
Phyllanthus urinaria
Sri Lanka
ND
I, E
[18]
Phytolaccaceae
Petiveria
a
l
l
i
a
c
e
a
#
Colombia, Nicaragua
Branch, leaf, root, whole plant
I, E
[27, 34, 35]
Pinaceae
Pinus
r
o
x
b
u
r
g
h
i
i
∗
Pakistan
Oil, resin, wood
E
[17, 53]
Piperaceae
Peperomia
e
l
s
a
n
a
#
Colombia
Whole plant
E
[35]
Peperomia pellucida
Nicaragua, Sri Lanka
Whole plant
I, E
[18, 27]
Piper amalago
Mexico, Nicaragua
Leaf, root
I
[27, 59]
Piper
a
r
b
o
r
e
u
m
∗
Colombia
Branch, leaf
E
[35]
Piper
a
u
r
i
t
u
m
#
Colombia, Nicaragua
Branch, leaf, stem, whole plant
I, E
[27, 34, 35, 44]
Piper betle
Sri Lanka
ND
I, E
[18]
Piper chuvya
Sri Lanka
ND
E
[18]
Piper confusionis
Peru
Leaf
E
[57]
Piper
c
o
r
u
s
c
a
n
s
#
Colombia
Branch, leaf, stem
I, E
[35]
Piper
h
i
s
p
i
d
u
m
#
Colombia
Branch, leaf, stem
I, E
[35]
Piper
l
o
n
g
i
v
i
l
l
o
s
u
m
#
Colombia
Whole plant
E
[35]
Piper
l
o
n
g
u
m
∗
Bangladesh, Sri Lanka
Flower, fruit, Latex, root
E
[18, 30]
Piper
m
a
r
g
i
n
a
t
u
m
#
Brazil, Colombia
Branch, leaf, root, stem
I, E
[35, 50]
Piper
m
u
l
t
i
p
l
i
n
e
r
v
i
u
m
#
Colombia
Branch, leaf, stem
I, E
[35]
Piper nigrum
Bangladesh, India, Sri Lanka
Floral bud, flower, fruit, root
I, E
[18, 20, 28, 32, 52]
Piper
p
e
l
t
a
t
u
m
#
Colombia, Nicaragua
Branch, leaf, stem, whole plant
I, E
[27, 35]
Piper
p
u
l
c
h
r
u
m
∗
Colombia
Branch, leaf, stem
I, E
[35]
Piper
r
e
t
i
c
u
l
a
t
u
m
#
Colombia
Branch, leaf, stem
I, E
[35]
Piper
t
r
i
c
u
s
p
e
#
Colombia
Branch, leaf, stem
E
[35]
Piper umbellatum
Sri Lanka
ND
I, E
[18]
Pittosporaceae
Pittosporum neelgherrense
India
Bark
I, E
[16]
Pittosporum tetraspermum
India
Bark
I
[26]
Plantaginaceae
Bacopa monnieri
Bangladesh, India, Sri Lanka
Leaf, root, whole plant
I
[18, 23, 32, 39, 41]
Plantago australis
Colombia
Whole plant
E
[34]
Plantago major
Colombia
Aerial parts, leaf
I, E
[44]
Scoparia
d
u
l
c
i
s
#
Colombia, Nicaragua
Aerial parts, branch, leaf, root, whole plant
I, E
[27, 34, 35, 44]
Platanaceae
Platanus orientalis
Pakistan
Bark
I, E
[17]
Plumbaginaceae
Plumbago indica
Sri Lanka
ND
I, E
[18]
Plumbago zeylanica
Bangladesh, India, Sri Lanka
Root
I, E
[18, 23, 32]
Poaceae
Chrysopogon zizanioides(syn. Vetiveria zizanioides)
India, Sri Lanka
Root
I, E
[16, 18]
Cymbopogon citratus
Colombia
Leaf
E
[34]
Cynodon dactylon
Bangladesh, India, Sri Lanka
Leaf, root, whole plant
E
[18, 19, 32]
Drynaria quercifolia
Sri Lanka
ND
I
[18]
Eleusine coracana
Sri Lanka
ND
I
[18]
Gynerium sagittatum
Nicaragua
Leaf, root
I
[27]
Heteropogon contortus
India, Sri Lanka
Root
I, E
[18, 38, 55]
Isachne globosa
Sri Lanka
ND
E
[18]
Oryza punctata
Sri Lanka
ND
I, E
[18]
Oryza sativa
Sri Lanka
ND
I
[18]
Pogonatherum paniceum
Sri Lanka
ND
E
[18]
Saccharum arundinaceum
Sri Lanka
ND
I
[18]
Saccharum officinarum
Colombia, Sri Lanka
Stem
I, E
[18, 34, 44]
Polygalaceae
Polygala abyssinica
Pakistan
Root
I
[17]
Polygala crotalarioides
India
Leaf, root
ND
[19]
Polygala paniculata
Brazil
Root
E
[47]
Polygala spectabilis
Brazil
Root
I, E
[47]
Polygonaceae
Persicaria barbata(syn.Polygonum barbatum)
India
Leaf
I, E
[38]
Persicaria chinensis(syn. Polygonum chinense)
Bangladesh
Leaf
E
[32]
Persicaria ferruginea(syn.Polygonum ferrugineum)
Colombia
Aerial parts
E
[34]
Persicaria glabra(syn.Polygonum glabrum)
India
Root
E
[25]
Polypodiaceae
Pleopeltis
p
e
r
c
u
s
s
a
∗
Colombia
Branch, leaf, stem
I, E
[35]
Pyrrosia piloselloides
Sri Lanka
ND
E
[18]
Pontederiaceae
Monochoria hastata
Sri Lanka
ND
I, E
[18]
Portulacaceae
Portulaca pilosa
Brazil
Leaf
I
[33]
Primulaceae
Aegiceras corniculatum
Sri Lanka
ND
E
[18]
Anagallis arvensis
Serbia
Aerial parts
ND
[58]
Ardisia humilis
Sri Lanka
ND
E
[18]
Maesa
l
a
n
c
e
o
l
a
t
a
∗
Kenya
Root
ND
[31]
Myrsine coriacea
Colombia
Whole plant
E
[34]
Pteridaceae
Acrostichum aureum
Nicaragua
Leaf, root
I, E
[27]
Adiantum capillus-veneris
Pakistan
Frond
E
[17]
Pellaea viridis
Kenya
Leaf
E
[31]
Ranunculaceae
Clematis brachiata(syn.Clematis triloba)
India
Root
E
[25]
Delphinium denudatum
India
Root
ND
[19]
Delphinium vestitum
India
Whole plant
ND
[19]
Rhamnaceae
Alphitonia incana
Papua New Guinea
Oil
E
[46]
Ziziphus jujuba(syn.Ziziphus mauritiana)
Sri Lanka
ND
E
[18]
Ziziphus oenoplia
India, Sri Lanka
Leaf
I, E
[18, 49]
Rhizophoraceae
Rhizophora mangle
Nicaragua
Bark
I, E
[27]
Rosaceae
Crataegus monogyna
Spain
Thorn
ND
[37]
Potentilla sundaica
India
Root, stem
ND
[19]
Prunus persica
Ethiopia
Leaf
I
[51]
Prunus walkeri
Sri Lanka
ND
E
[18]
Pyrus communis
Pakistan
Fruit, leaf
I
[17]
Sanguisorba officinalis
Serbia
Rhizome
ND
[58]
Rubiaceae
Catunaregam spinosa(syn. Randia dumetorum)
India
Root
I
[23]
Ceriscoides turgida(syn. Gardenia turgida)
India
Bark, root
I
[24, 38]
Chiococca alba
Brazil, Nicaragua
Leaf, root
I
[27, 47]
Clausena dentata
Sri Lanka
ND
E
[18]
Gonzalagunia
p
a
n
a
m
e
n
s
i
s
∗
Colombia
Branch, leaf, stem
I, E
[35]
Hamelia axillaris
Nicaragua
Leaf, whole plant
I, E
[27]
Hamelia barbata
Nicaragua
Leaf, whole plant
I, E
[27]
Hamelia patens
Nicaragua
Leaf, whole plant
I, E
[27]
Hamelia rovirosae
Nicaragua
Flower, leaf, stem
I, E
[27]
Hedyotis scandens
Bangladesh
Leaf, stem
E
[32]
Ixora coccinea
Sri Lanka
ND
I, E
[18]
Ixora cuneifolia
Bangladesh
Bark
E
[32]
Ixora pavetta(syn.Ixora arborea)
India
Leaf, rood, seed
ND
[19]
Mitragyna parvifolia
India
Bark, stem
I, E
[38, 63]
Morinda angustifolia
Bangladesh
Leaf
I
[32]
Morinda citrifolia
Bangladesh
Root
ND
[30]
Morinda coreia
Sri Lanka
ND
I, E
[18]
Morinda persicifolia
Bangladesh
Leaf
E
[32]
Mussaenda frondosa
Sri Lanka
ND
I
[18]
Mussaenda roxburghii
Bangladesh
Leaf
E
[32]
Nauclea orientalis
Sri Lanka
ND
E
[18]
Neonauclea purpurea(syn. Anthocephalus chinensis)
Bangladesh
Bark, leaf
ND
[30]
Oldenlandia diffusa
India
Whole plant
E
[20]
Oldenlandia umbellata
India
Leaf, root
E
[20]
Ophiorrhiza
m
u
n
g
o
s
∗
India
Root
I
[16, 20]
Paederia foetida
Sri Lanka
ND
I, E
[18]
Palicourea croceoides
Colombia
Bark
I
[34]
Pavetta indica
Sri Lanka
ND
I, E
[18]
Psychotria elata
Nicaragua
Flower, leaf, root, stem, whole plant
I, E
[27]
Psychotria flavida
India
Root
I
[63]
Psychotria
p
o
e
p
p
i
g
i
a
n
a
#
Colombia, Nicaragua, Sri Lanka
Branch, leaf, stem, whole plant
I, E
[18, 27, 35]
Randia
a
c
u
l
e
a
t
a
∗
Mexico
Fruit, whole plant
I
[59, 78]
Rubia
c
o
r
d
i
f
o
l
i
a
∗
Nepal, Pakistan
Leaf, root, stem
I
[17, 69]
Rubia manjith
India
Root, stem
ND
[19]
Spermacoce remota(syn.Borreria assurgens)
Nicaragua
Leaf, root
I, E
[27]
Tamilnadia uliginosa
Sri Lanka
ND
I
[18]
Wendlandia exserta
India
Root
I
[49]
Rutaceae
Acronychia pedunculata
Sri Lanka
ND
E
[18]
Aegle marmelos
Bangladesh, India, Sri Lanka
Bark, whole plant
I, E
[18, 20, 30, 32, 41]
Atalantia ceylanica
Sri Lanka
ND
I, E
[18]
Citrus aurantiifolia
Sri Lanka
ND
I, E
[18]
Citrus aurantium
Sri Lanka
ND
I, E
[18]
Citrus japonica(syn. Citrus madurensis)
Sri Lanka
ND
I, E
[18]
Citrus
l
i
m
o
n
∗
Colombia, India, Sri Lanka
Fruit, leaf, root
I, E
[18, 20, 28, 34, 35]
Citrus maxima(syn. Citrus grandis)
Sri Lanka
ND
I, E
[18]
Glycosmis pentaphylla
India
Leaf
I, E
[16]
Limonia acidissima(syn.Feronia limonia)
India, Sri Lanka
Root
I
[18, 20]
Murraya koenigii
India, Sri Lanka
Bark, leaf
I, E
[18, 28]
Murraya
p
a
n
i
c
u
l
a
t
a
∗
Sri Lanka
ND
E
[18]
Naringi crenulata
India
Fruit
ND
[19]
Pamburus missionis
Sri Lanka
ND
E
[18]
Ruta chalepensis
Colombia
Whole plant
E
[34]
Toddalia asiatica
India, Sri Lanka
Root
I, E
[18, 63]
Salicaceae
Casearia
g
r
a
n
d
i
f
l
o
r
a
∗
ND
Bark, leaf
ND
[79]
Casearia nigrescens(syn.Casearia elliptica)
India
Bark, leaf
ND
[19]
Casearia
s
y
l
v
e
s
t
r
i
s
∗
Brazil
Leaf, whole plant
ND
[47, 79]
Casearia tomentosa
India
Bark, root
I, E
[49, 79]
Flacourtia indica
Bangladesh
Leaf
E
[32]
Santalaceae
Santalum album
Sri Lanka
ND
E
[18]
Sapindaceae
Allophylus cobbe
Sri Lanka
ND
I, E
[18]
Cardiospermum halicacabum
India, Sri Lanka
Leaf
I, E
[18, 28]
Dodonaea viscosa
India
Leaf
E
[28]
Harpullia arborea
Sri Lanka
ND
I, E
[18]
Sapindus emarginatus
India
Bark
I
[20]
Sapindus mukorossi
India, Pakistan
Fruit, leaf, root, seed
E
[17, 25]
Sapotaceae
Madhuca longifolia(syn.Madhuca indica)
India, Sri Lanka
Bark, fruit, leaf, nut, root, seed
I, E
[18, 20, 23, 25, 32, 38]
Manilkara zapota
Mexico
Root
ND
[59]
Mimusops elengi
Sri Lanka
ND
I
[18]
Scrophulariaceae
Verbascum thapsus
India
Leaf
ND
[66]
Selaginellaceae
Selaginella
a
r
t
i
c
u
l
a
t
a
#
Colombia
Whole plant
I, E
[35]
Simaroubaceae
Ailanthus excelsa
India
Bark
I
[38]
Quassia
a
m
a
r
a
#
Colombia, Nicaragua
Root, stem, whole plant
I, E
[27, 35]
Quassia indica
Sri Lanka
ND
I
[18]
Simaba
c
e
d
r
o
n
#
Colombia
Seed, whole plant
I, E
[34, 35, 44]
Siparunaceae
Siparuna gesnerioides
Colombia
Leaf, root
I
[34, 44]
Siparuna
t
h
e
c
a
p
h
o
r
a
∗
Colombia
Branch, leaf, stem
I, E
[35]
Smilacaceae
Smilax regelii
Nicaragua
Root
I
[27]
Smilax spinosa
Nicaragua
Root
I
[27]
Solanaceae
Atropa acuminata
Pakistan
Leaf, root
E
[17]
Capsicum annuum(syn.Capsicum frutescens)∗
Bangladesh, Colombia, India, Sri Lanka
Fruit, root
I, E
[18, 19, 25, 32, 34, 35]
Datura metel
Bangladesh, Colombia, India, Sri Lanka
Bark, flower, fruit, leaf, root, seed
I, E
[18, 22, 23, 25, 28, 30, 34]
Datura
s
t
r
a
m
o
n
i
u
m
#
India
Root
I, E
[38]
Lycopersicon esculentum
Colombia
Leaf, stem, whole plant
E
[34]
Nicotiana tabacum
Colombia, India, Nicaragua
Leaf
I, E
[20, 27, 44]
Solanum
a
l
l
o
p
h
y
l
l
u
m
#
Colombia
Branch, leaf, stem
I, E
[35]
Solanum americanum(syn.Solanum nigrum)
Colombia, India, Sri Lanka
Fruit, leaf, whole plant
I, E
[18, 25, 34, 38]
Solanum capsicoides
Bangladesh
Seed
ND
[30]
Solanum incanum
Kenya
Fruit, stem
E
[31]
Solanum melongena
Sri Lanka
ND
I
[18]
Solanum
n
u
d
u
m
#
Colombia
Branch, fruit, leaf, stem
I, E
[35]
Solanum ochraceo-ferrugineum
Mexico
Whole plant
ND
[59]
Solanum torvum
Bangladesh, India, Nicaragua
Flower, leaf, root
I, E
[20, 27, 30, 32]
Solanum verbascifolium
Mexico
Whole plant
ND
[59]
Solanum virginianum(syn. Solanum xanthocarpum)
India, Sri Lanka
Root
I
[18, 41]
Withania somnifera
Bangladesh, India, Sri Lanka
Root
I, E
[18, 22, 32]
Sterculiaceae
Byttneria pilosa
Bangladesh
Leaf, stem
E
[32]
Symplocaceae
Symplocos cochinchinensis
Sri Lanka
ND
E
[18]
Symplocos racemosa
Sri Lanka
ND
I, E
[18]
Talinaceae
Talinum paniculatum
Sri Lanka
ND
E
[18]
Thymelaeaceae
Daphne papyracea
Pakistan
Leaf, root, stem
ND
[17]
Triuridaceae
Sciaphila purpurea
Colombia
Whole plant
I, E
[35]
Urticaceae
Boehmeria nivea
Sri Lanka
ND
E
[18]
Cecropia obtusifolia
Nicaragua
Leaf
I
[27]
Cecropia peltata
Bangladesh, Nicaragua
Leaf, whole plant
I
[27, 30]
Girardinia diversifolia
Sri Lanka
ND
E
[18]
Pouzolzia zeylanica(syn.Pouzolzia indica)
Bangladesh, India
Leaf, whole plant
E
[32, 42]
Verbenaceae
Aloysia triphylla
Colombia
Aerial parts
E
[34]
Lantana camara
Bangladesh, Colombia, India, Sri Lanka
Flower, leaf, root, stem
I, E
[18, 32, 34, 39, 41]
Lippia alba
Colombia
Aerial parts, leaf, whole plant
I, E
[34]
Lippia grandis
Brazil
Leaf
I
[33]
Stachytarpheta cayennensis
Colombia
Whole plant
E
[34]
Verbena litoralis
Colombia
Fruit peel, leaf, root, seed
E
[34]
Verbena officinalis
Pakistan
Root, whole plant
E
[17, 53]
Vitaceae
Ampelocissus latifolia
India
Root
I
[49]
Cayratia pedata(syn. Cissus pedata)
Bangladesh
Leaf
ND
[30]
Cayratia trifolia(syn. Vitis trifolia)
Bangladesh, India
Leaf, root
I, E
[36, 38]
Cissus adnata
Bangladesh
Leaf
E
[32]
Cissus javana
Bangladesh
Leaf, stem
E
[32]
Cissus
q
u
a
d
r
a
n
g
u
l
a
r
i
s
#
Sri Lanka
ND
E
[18]
Leea indica
Sri Lanka
ND
E
[18]
Vitis heyneana(syn.Vitis lanata)
Bangladesh
Leaf
E
[32]
Xanthorrhoeaceae
Aloe harlana
Ethiopia
Leaf
I
[51]
Aloe littoralis
Pakistan
Whole plant
E
[17]
Aloe vera
Nicaragua, Sri Lanka
Leaf
I, E
[18, 27]
Zingiberaceae
Alpinia calcarata
Sri Lanka
ND
I, E
[18]
Alpinia galanga
Sri Lanka
ND
I
[18]
Alpinia nigra
Sri Lanka
ND
E
[18]
Alpinia purpurata
Colombia
Leaf
E
[44]
Curcuma angustifolia
India
Rhizome
E
[28]
Curcuma
l
o
n
g
a
∗
Bangladesh, India, Sri Lanka
Rhizome
I, E
[16, 18, 20, 30]
Elettaria cardamomum
Sri Lanka
ND
I
[18]
Globba marantina(syn.Globba bulbifera)
India
Rhizome
I
[49]
Hedychium coronarium
Colombia, Nicaragua
Root, whole plant
E
[27, 34]
Renealmia
a
l
p
i
n
i
a
∗
Colombia
Rhizome, stem
I, E
[34, 35]
Renealmia thyrsoidea
Colombia
Leaf, stem
I
[34]
Zingiber
o
f
f
i
c
i
n
a
l
e
∗
Ecuador, Nicaragua, Sri Lanka
Rhizome, root
I, E
[18, 27, 61]
Zygophyllaceae
Balanites
a
e
g
y
p
t
i
a
c
a
#
India
Bark, fruit
E
[38]
In parentheses is the synonym used in the original work; out of the parentheses is the accepted name (in case of more than one paper treating the same species with different names); ND = information not described in the work; I = internal use; E = external use.∗Species evaluated on antiophidic activities in previous studies (see Tables 2–8) showing good inhibitory potential against venom induced local effects. #Species evaluated on antiophidic activities in previous studies, however, with poor inhibition potential against venom induced local effects.Table 2
List of medicinal plants with inhibitory potential against local effects induced byNajasnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Acanthaceae
Andrographis stenophylla
Leaf
N. naja
—
Hemorrhage
[83]
Amaranthaceae
Pupalia lappacea
Herbal
N. nigricollis
SVH
—
[82]
Amaryllidaceae
Allium
c
e
p
a
#
Bulb
N. n. karachiensis
PLA2
—
[84]
Allium
s
a
t
i
v
u
m
#
Bulb
N. n. karachiensis
PLA2
—
[84]
Anacardiaceae
Lannea acida
Cortex
N. nigricollis
SVH
—
[82]
Pistacia chinensissubsp.integerrima#
Gall
N. n. karachiensis
PLA2
—
[84]
Sclerocarya birrea
Cortex
N. nigricollis
SVH
—
[82]
Spondias
m
o
m
b
i
n
#
Cortex, radix
N. nigricollis
SVH
—
[82]
Annonaceae
Annona
s
e
n
e
g
a
l
e
n
s
i
s
#
Cortex
N. nigricollis
SVH
—
[82]
Apiaceae
Cuminum cyminum
Seed
N. n. karachiensis
PLA2
—
[84]
Apocynaceae
Acokanthera oppositifolia
Radix
N. nigricollis
SVH
—
Calotropis
p
r
o
c
e
r
a
#
Flower, latex
N. n. karachiensis
PLA2
—
[84]
Strophanthus sarmentosus
Folium
N. nigricollis
SVH
—
[82]
Strophanthus speciosus
Radix
N. nigricollis
SVH
—
[82]
Tylophora
i
n
d
i
c
a
#
Leaf, root
N. naja
PLA2
Hemorrhage
[85]
Araceae
Colocasia esculenta
Tuber
N. nigricollis
SVH
—
[82]
Araliaceae
Polyscias fulva
Cortex
N. nigricollis
SVH
—
[82]
Aristolochiaceae
Aristolochia
b
r
a
c
t
e
o
l
a
t
a
#
Leaf, root
N. naja
PLA2
Hemorrhage
[85]
Asteraceae
Callilepis laureola
Radix
N. nigricollis
SVH
—
[82]
Bignoniaceae
Kigelia africana
Cortex, folium
N. nigricollis
SVH
—
[82]
Tecoma stans(syn. Stenolobium stans)
Root
N. n. karachiensis
PLA2
—
[84]
Bixaceae
Cochlospermum tinctorium
Radix
N. nigricollis
SVH
—
[82]
Boraginaceae
Cordia macleodii
Bark
N. naja
—
Edema, hemorrhage∗, necrosis∗
[86]
Trichodesma
i
n
d
i
c
u
m
#
Whole plant
N. n. karachiensis
PLA2
—
[84]
Capparaceae
Capparis tomentosa
Radix
N. nigricollis
SVH
—
[82]
Colchicaceae
Gloriosa
s
u
p
e
r
b
a
#
Radix
N. nigricollis
SVH
—
[82]
Combretaceae
Combretum
m
o
l
l
e
#
Folium
N. nigricollis
SVH
—
[82]
Guiera senegalensis
Radix
N. nigricollis
SVH
—
[82]
Terminalia
a
r
j
u
n
a
#
Bark
N. n. karachiensis
PLA2
—
[84]
Convolvulaceae
Ipomoea rubens
Seed
N. nigricollis
SVH
—
[82]
Cucurbitaceae
Citrullus
c
o
l
o
c
y
n
t
h
i
s
#
Fruit
N. n. karachiensis
PLA2
—
[84]
Luffa cylindrica(syn.Luffa aegyptiaca)
Leaf
N. nigricollis
Proteolytic
—
[87]
Momordica
c
h
a
r
a
n
t
i
a
#
Fruit
N. n. karachiensis
PLA2
—
[84]
Ebenaceae
Diospyros mespiliformis
Cortex
N. nigricollis
SVH
—
[82]
Euphorbiaceae
Alchornea laxiflora
Cortex
N. nigricollis
SVH
—
[82]
Clutia cordata
Radix
N. nigricollis
SVH
—
[82]
Euphorbia
h
i
r
t
a
#
Whole plant
N. naja
PLA2, proteolytic, SVH
Edema∗
[88]
Jatropha
c
u
r
c
a
s
#
Leaf, root, stem
N. naja
PLA2
—
[89]
Jatropha
g
o
s
s
y
p
i
i
f
o
l
i
a
#
Leaf, root, stem
N. naja
PLA2
—
[89]
Manihot foetida(syn.Jatropha foetida)
Leaf, stem
N. naja
PLA2
—
[89]
Fabaceae
Abrus
p
r
e
c
a
t
o
r
i
u
s
#
Radix
N. nigricollis
SVH
—
[82]
Argyrolobium stipulaceum
Radix
N. nigricollis
SVH
—
[82]
Bauhinia thonningii
Cortex, radix
N. nigricollis
SVH
—
[82]
Bauhinia
v
a
r
i
e
g
a
t
a
#
Root
N. n. karachiensis
PLA2
—
[84]
Bobgunnia madagascariensis(syn.Swartzia madagascariensis)
Folium, radix
N. nigricollis
SVH
—
[82]
Burkea africana
Cortex
N. nigricollis
SVH
—
[82]
Cullen corylifolium(syn.Psoralea corylifolia)
Seed
N. n. karachiensis
PLA2
—
[84]
Dichrostachys cinerea
Folium
N. nigricollis
SVH
—
[82]
Entada africana
Radix
N. nigricollis
SVH
—
[82]
Mimosa
p
u
d
i
c
a
#
Root
N. kaouthia, N. naja
PLA2, proteolytic, SVH
Edema∗, myotoxicity∗
[90–92]
Parkia biglobosa
Cortex, stem bark
N. nigricollis
Cytotoxicity against muscle cells, SVH
[82, 93]
Stylosanthes erecta
Folium
N. nigricollis
SVH
—
[82]
Tamarindus
i
n
d
i
c
a
#
Folium, radix
N. nigricollis
SVH
—
[82]
Gentianaceae
Enicostema axillare(syn. Enicostema hyssopifolium)#
Whole plant
N. n. karachiensis
PLA2
—
[84]
Hypericaceae
Psorospermum corymbiferum
Cortex, radix
N. nigricollis
SVH
—
[82]
Lamiaceae
Leucas
a
s
p
e
r
a
#
Leaf, root, whole plant
N. naja
PLA2, proteolytic, SVH
Hemorrhage
[85, 94]
Leucas cephalotes(syn.Leucas capitata)#
Whole plant
N. n. karachiensis
PLA2
—
[84]
Leucas martinicensis
ND
N. nigricollis
SVH
—
[82]
Ocimum tenuiflorum(syn.Ocimum sanctum)#
Whole plant
N. n. karachiensis
PLA2
—
[84]
Rotheca myricoides(syn. Clerodendrum myricoides)
Cortex
N. nigricollis
SVH
—
[82]
Teucrium kraussii
Aerial parts, cortex
N. nigricollis
SVH
—
[82]
Volkameria glabra(syn.Clerodendrum glabrum)
Radix
N. nigricollis
SVH
—
[82]
Lauraceae
Cassytha filiformis
Herbal
N. nigricollis
SVH
—
[82]
Loganiaceae
Strychnos innocua
Folium
N. nigricollis
SVH
—
[82]
Strychnos
nux
-vomica
#
Seed
N. kaouthia
PLA2
—
[95]
Malvaceae
Althaea officinalis
Root
N. n. karachiensis
PLA2
—
[84]
Dombeya quinqueseta
Cortex
N. nigricollis
SVH
—
[82]
Grewia mollis
Cortex, folium, radix
N. nigricollis
SVH
—
[82]
Sterculia setigera
Cortex
N. nigricollis
SVH
—
[82]
Waltheria indica
Radix
N. nigricollis
SVH
—
[82]
Menispermaceae
Cissampelos mucronata
Herbal
N. nigricollis
SVH
—
[82]
Moraceae
Ficus platyphylla
Folium
N. nigricollis
SVH
—
[82]
Olacaceae
Ximenia americana
Folium
N. nigricollis
SVH
—
[82]
Pedaliaceae
Ceratotheca sesamoides
Herbal
N. nigricollis
SVH
—
[82]
Peraceae
Clutia pulchella
Radix
N. nigricollis
SVH
—
[82]
Phyllanthaceae
Flueggea virosa(syn.Securinega virosa)
Radix
N. nigricollis
SVH
—
[82]
Pinaceae
Cedrus deodara
Bark
N. n. karachiensis
PLA2
—
[84]
Pinus
r
o
x
b
u
r
g
h
i
i
#
Oleoresin
N. n. karachiensis
PLA2
—
[84]
Poaceae
Cymbopogon schoenanthus
Radix
N. nigricollis
SVH
—
[82]
Primulaceae
Maesa
l
a
n
c
e
o
l
a
t
a
#
Cortex
N. nigricollis
SVH
—
[82]
Rhamnaceae
Ziziphus mucronata
Radix
N. nigricollis
SVH
—
[82]
Ziziphus spina-christi
Cortex
N. nigricollis
SVH
—
[82]
Rubiaceae
Crossopteryx febrifuga
Cortex
N. nigricollis
SVH
—
[82]
Pentanisia prunelloides
Radix
N. nigricollis
SVH
—
[82]
Pentas zanzibarica
Folium
N. nigricollis
SVH
—
[82]
Rubia
c
o
r
d
i
f
o
l
i
a
#
Stem
Rutaceae
Citrus
l
i
m
o
n
#
Fruit
N. n. karachiensis
PLA2
—
[84]
Zanthoxylum capense
Radix
N. nigricollis
SVH
—
[82]
Sapindaceae
Paullinia pinnata
Folium
N. nigricollis
SVH
—
[82]
Sapindus mukorossi
Fruit
N. n. karachiensis
PLA2
—
[84]
Solanaceae
Nicotiana rustica
Leaf
N. nigricollis
Proteolytic
—
[87]
Schwenckia americana
Folium
N. nigricollis
SVH
—
[82]
Thymelaeaceae
Gnidia anthylloides
Radix
N. nigricollis
SVH
—
[82]
Gnidia kraussiana
Radix
N. nigricollis
SVH
—
[82]
Gnidia splendens
Radix
N. nigricollis
SVH
—
[82]
Verbenaceae
Lantana trifolia
Cortex
N. nigricollis
SVH
—
[82]
Vitaceae
Cissus populnea
Stem
N. nigricollis
SVH
—
[82]
Zingiberaceae
Zingiber
o
f
f
i
c
i
n
a
l
e
#
Rhizome
N. n. karachiensis
PLA2
—
[84]
Zygophyllaceae
Fagonia cretica
Leaf, stem
N. n. karachiensis
PLA2
—
[84]
ND = information not described in the work; PLA2 = snake venom phospholipase A2; SVH = snake venom hyaluronidase. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details).Table 3
List of medicinal plants with inhibitory potential against local effects induced byBothropssnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Acanthaceae
Justicia
p
e
c
t
o
r
a
l
i
s
#
Leaf
B. jararaca
—
Hemorrhage∗
[33]
Amaranthaceae
Blutaparon portulacoides
Aerial parts
B. jararacussu
—
Edema
[109]
Anacardiaceae
Anacardium excelsum
Leaf, twig
B. asper
PLA2
—
[111]
Annonaceae
Ephedranthus columbianus
Leaf, twig
B. asper
PLA2
—
[111]
Sapranthus isae
Leaf, twig
B. asper
PLA2
—
[111]
Apocynaceae
Allamanda
c
a
t
h
a
r
t
i
c
a
#
Branch, leaf, stem
B. atrox
—
Hemorrhage∗
[124]
Fernaldia pandurata(syn.Mandevilla velutina)
Leaf, stem, subterranean system
B. alternatus, B. jararacussu, B. moojeni, B. pirajai
PLA2
Edema∗, hemorrhage∗, myotoxicity∗
[125]
Tabernaemontana catharinensis
Root bark
B. jararacussu
Myotoxicity
Myotoxicity∗∗
[126]
Araceae
Dracontium
c
r
o
a
t
i
i
#
Rhizome
B. asper
—
Edema∗
[127]
Philodendron
m
e
g
a
l
o
p
h
y
l
l
u
m
#
Liana, vine
B. atrox, B. jararaca
PLA2
Edema∗, hemorrhage∗
[33, 128]
Philodendron
t
r
i
p
a
r
t
i
t
u
m
#
Branch, leaf
B. atrox
—
Hemorrhage∗
[124]
Asteraceae
Chaptalia nutans
Leaf
B. asper
—
Edema
[129]
Eclipta prostrata(syn.Eclipta alba)#
Aerial parts
B. jararaca, B. jararacussu
Myotoxicity
Myotoxicity
[130]
Mikania glomerata
Leaf
B. jararaca
—
Edema∗, hemorrhage∗, peritonitis
[131, 132]
Neurolaena
l
o
b
a
t
a
#
Branch, leaf, stem
B. atrox
—
Hemorrhage∗
[124]
Pseudelephantopus
s
p
i
c
a
t
u
s
#
Whole plant
B. atrox
—
Hemorrhage∗
[124]
Bignoniaceae
Fridericia chica(syn.Arrabidaea chica)
Leaf
B. atrox
—
Edema
[133]
Tabebuia aurea
Stem bark
B. neuwiedi
H2O2 production by peritoneal macrophages
Edema, hemorrhage∗, myonecrosis∗, peritonitis
[110]
Tabebuia
r
o
s
e
a
#
Stem bark
B. asper, B. atrox
PLA2
Edema∗∗, Hemorrhage∗
[124, 127, 134]
Bixaceae
Bixa
o
r
e
l
l
a
n
a
#
Branch, leaf
B. asper, B. atrox
PLA2
Edema∗∗, hemorrhage∗
[124, 127, 134]
Boraginaceae
Cordia verbenacea
Leaf
B. jararacussu
—
Edema∗
[135]
Clusiaceae
Clusia fluminensis
Fruit
B. jararaca
Proteolytic
Hemorrhage∗∗
[136]
Combretaceae
Combretum leprosum
Root
B. jararaca, B. jararacussu
Collagenase, myotoxicity, PLA2, proteolytic
Edema, hemorrhage∗, myotoxicity∗∗
[99]
Connaraceae
Connarus
f
a
v
o
s
u
s
#
Bark
B. atrox, B. jararaca
PLA2, proteolytic
Edema∗, hemorrhage∗∗∗
[33, 115, 128]
Costaceae
Costus
l
a
s
i
u
s
#
ND
B. atrox
PLA2
—
[134]
Costus spicatus
Leaf
B. atrox
—
Edema∗, hyperalgesia∗
[137]
Crassulaceae
Bryophyllum pinnatum(syn.Kalanchoe pinnata)#
Leaf
B. jararaca
PLA2
Edema, hemorrhage
[138]
Kalanchoe laciniata(syn.Kalanchoe brasiliensis)#
Leaf
B. jararaca
PLA2
Edema, hemorrhage∗∗∗
[33, 138]
Dicranaceae
Dicranum frigidum
Whole plant
B. asper
PLA2
—
[111]
Dilleniaceae
Davilla elliptica
Leaf
B. jararaca
—
Hemorrhage∗
[139]
Euphorbiaceae
Croton urucurana
Stem bark
B. jararaca
—
Hemorrhage∗
[140]
Hevea nitida
Leaf, twig
B. asper
PLA2
—
[111]
Jatropha
g
o
s
s
y
p
i
i
f
o
l
i
a
#
Leaf
B. erythromelas, B. jararaca
PLA2, proteolytic, SVH
Edema, hemorrhage, myotoxicity
[96, 119]
Jatropha
m
o
l
l
i
s
s
i
m
a
#
Leaf
B. erythromelas, B. jararaca
—
Edema, hemorrhage, myotoxicity, peritonitis
[141]
Fabaceae
Abarema cochliacarpos
Stem bark
B. leucurus
—
Decreased motor function, edema, hyperalgesia, myotoxicity
[101]
Brownea ariza
Bark, leaf
B. asper
PLA2, proteolytic
—
[142]
Brownea
rosa
-de-monte
#
Leaf, stem bark
B. asper, B. atrox
PLA2, proteolytic
Edema∗∗∗, hemorrhage∗∗∗
[124, 127, 134, 143]
Cassia
f
i
s
t
u
l
a
#
Seed
B. jararaca
—
Hemorrhage∗
[33]
Dipteryx alata
Bark
B. jararacussu
Myotoxicity
—
[144]
Pentaclethra
m
a
c
r
o
l
o
b
a
#
Bark
B. alternatus, B. asper, B. atrox, B. jararaca, B. jararacussu, B. moojeni, B. neuwiedi, B. pirajai
PLA2
Edema∗, hemorrhage∗, myotoxicity∗
[145]
Plathymenia
r
e
t
i
c
u
l
a
t
a
#
Bark
B. atrox, B. jararaca
PLA2
Edema∗, hemorrhage∗
[33, 128]
Schizolobium parahyba
Leaf
B. alternatus, B. moojeni, B. pauloensis
PLA2
Hemorrhage∗, myotoxicity∗∗∗
[146, 147]
Senna
d
a
r
i
e
n
s
i
s
#
Whole plant
B. atrox
PLA2
Hemorrhage∗
[124, 134]
Heliconiaceae
Heliconia
c
u
r
t
i
s
p
a
t
h
a
#
Rhizome
B. asper, B. atrox
PLA2, proteolytic
Edema∗∗∗, hemorrhage∗
[124, 127, 134, 148]
Heliconia latispatha
Rhizome
B. asper
PLA2, proteolytic
—
[148]
Heliconia wagneriana
Rhizome
B. asper
PLA2, proteolytic
—
[148]
Hymenophyllaceae
Trichomanes elegans#
Whole plant
B. asper, B. atrox
PLA2
Edema∗∗∗, hemorrhage∗
[124, 127, 134]
Hypericaceae
Hypericum brasiliense
Whole plant
B. jararaca
Proteolytic
Edema∗, hemorrhage∗
[149]
Icacinaceae
Casimirella ampla(syn.Humirianthera ampla)
Root
B. atrox, B. jararaca, B. jararacussu
Myotoxicity, PLA2, proteolytic
Edema∗∗∗, hemorrhage∗∗, myotoxicity∗
[102]
Lamiaceae
Marsypianthes
c
h
a
m
a
e
d
r
y
s
#
Inflorescence, leaf
B. atrox
PLA2
Peritonitis
[108]
Peltodon radicans
Flower, leaf, stem
B. atrox
—
Edema
[150]
Lauraceae
Aniba parviflora(syn. Aniba fragrans)#
Bark, leaf
B. atrox, B. jararaca
PLA2
Edema∗, hemorrhage∗
[33, 128]
Loasaceae
Nasa speciosa (syn. Loasa speciosa)
Leaf
B. asper
—
Edema
[129]
Loganiaceae
Strychnos pseudoquina
Leaf
B. jararaca
—
Hemorrhage∗
[139]
Strychnos
x
i
n
g
u
e
n
s
i
s
#
ND
B. atrox
PLA2
—
[134]
Loranthaceae
Struthanthus
o
r
b
i
c
u
l
a
r
i
s
#
Branch, leaf
B. asper, B. atrox
PLA2
Edema∗∗, hemorrhage∗
[124, 127, 134]
Magnoliaceae
Magnolia espinalii(syn.Talauma espinalii)
Leaf, twig
B. asper
PLA2
—
[111]
Magnolia guatapensis(syn.Dugandiodendron guatapense)
Leaf, twig
B. asper
PLA2
—
[111]
Magnolia hernandezii(syn.Talauma hernandezii)
Leaf, twig
B. asper
PLA2
—
[111]
Magnolia yarumalensis(syn.Dugandiodendron yarumalense)
Leaf, twig
B. asper
PLA2
—
[111]
Malpighiaceae
Byrsonima crassa
Leaf
B. jararaca
—
Hemorrhage∗
Malvaceae
Pachira glabra (syn. Bombacopsis glabra)
Root bark
B. pauloensis
—
Hemorrhage
[151]
Melastomataceae
Bellucia
d
i
c
h
o
t
o
m
a
#
Bark
B. atrox, B. jararaca
PLA2
Edema∗∗∗, hemorrhage∗∗
[33, 98, 128, 152]
Mouriri pusa
Leaf
B. jararaca
—
Hemorrhage∗
[139]
Meliaceae
Carapa guianensis
Leaf, twig
B. asper
PLA2
—
[111]
Cedrela odorata
Leaf, twig
B. asper
PLA2
—
[111]
Swietenia humilis
Leaf, twig
B. asper
PLA2
—
[111]
Swietenia macrophylla
Leaf, twig
B. asper
PLA2
—
[111]
Swietenia mahagoni
Leaf, twig
B. asper
PLA2
—
[111]
Menispermaceae
Cissampelos
p
a
r
e
i
r
a
#
Leaf
B. asper
—
Hemorrhage∗
[153]
Moraceae
Brosimum guianense
Leaf
B. atrox
—
Hemorrhage∗, pain∗
[154]
Castilla
e
l
a
s
t
i
c
a
#
Branch, leaf, stem
B. atrox
—
Hemorrhage∗
[124]
Ficus
n
y
m
p
h
a
e
i
f
o
l
i
a
#
Branch, leaf, stem
B. asper, B. atrox
—
Edema∗∗, hemorrhage∗
[124, 127]
Musaceae
Musa × paradisíac
a
#
Exudate
B. jararacussu
PLA2
Hemorrhage∗∗, myonecrosis∗∗
[155]
Myrtaceae
Myrcia guianensis
Leaf
B. jararaca
PLA2
Hemorrhage∗
[156]
Passifloraceae
Passiflora
q
u
a
d
r
a
n
g
u
l
a
r
i
s
#
Branch, leaf
B. atrox
—
Hemorrhage∗
[124]
Piperaceae
Piper
a
r
b
o
r
e
u
m
#
Branch, leaf
B. atrox
PLA2
Hemorrhage∗
[124, 134]
Piper
p
u
l
c
h
r
u
m
#
Leaf, branch, stem
B. atrox
—
Hemorrhage∗
[124]
Polypodiaceae
Pleopeltis
p
e
r
c
u
s
s
a
#
Branch, leaf, stem, whole plant
B. asper, B. atrox
PLA2, proteolytic
Edema∗∗, hemorrhage∗
[124, 127, 134]
Rubiaceae
Gonzalagunia
p
a
n
a
m
e
n
s
i
s
#
Branch, leaf, stem
B. asper, B. atrox
PLA2
Edema∗∗, hemorrhage∗
[124, 127, 134]
Randia
a
c
u
l
e
a
t
a
#
Fruit
B. asper
—
Myotoxicity
[78]
Uncaria tomentosa
Root
B. asper
—
Edema
[129]
Rutaceae
Citrus
l
i
m
o
n
#
Ripe fruit
B. asper, B. atrox
—
Edema∗∗∗, hemorrhage∗
[124, 127]
Murraya
p
a
n
i
c
u
l
a
t
a
#
Leaf, twig
B. asper
PLA2
—
[111]
Salicaceae
Casearia
g
r
a
n
d
i
f
l
o
r
a
#
Leaf
B. moojeni, B. neuwiedi
PLA2
Myotoxicity∗
[157]
Casearia
s
y
l
v
e
s
t
r
i
s
#
Leaf
B. asper, B. jararacussu, B. moojeni, B. neuwiedi, B. pirajai
Myonecrosis, neuromuscular blockade
Edema∗, hemorrhage∗, myotoxicity∗
[158–160]
Sapindaceae
Billia hippocastanum
Leaf, twig
B. asper
PLA2
—
[111]
Cupania americana
Leaf, twig
B. asper
PLA2
—
[111]
Sapindus saponaria
In vitro cultivated callus, leaf, twig
B. alternatus, B. asper, B. jararacussu, B. moojeni
PLA2
Hemorrhage∗
[111, 161]
Serjania erecta
Aerial parts
B. jararacussu
PLA2
Edema∗, hemorrhage∗, myotoxicity∗
[162]
Siparunaceae
Siparuna
t
h
e
c
a
p
h
o
r
a
#
Branch, leaf, stem
B. atrox
—
Hemorrhage∗
[124]
Solanaceae
Capsicum annuum(syn.Capsicum frutescens)#
Ripe fruit
B. atrox
—
Hemorrhage∗
[124]
Urticaceae
Urera baccifera
Leaf
B. asper
—
Edema
[129]
Velloziaceae
Vellozia squamata(syn.Vellozia flavicans)
Leaf
B. jararacussu
Neuromuscular blockade and cell damage
—
[163]
Zingiberaceae
Curcuma
l
o
n
g
a
#
Rhizome
B. alternatus
—
Edema, hemorrhage, necrosis
[164]
Renealmia
a
l
p
i
n
i
a
#
Leaf, rhizome
B. asper, B. atrox
PLA2, proteolytic
Edema∗∗, hemorrhage
[107, 127, 134, 165, 166]
ND = information not described in the work; PLA2 = snake venom phospholipase A2; H2O2: hydrogen peroxide. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details). ∗∗Active in preincubation tests but inactive or only poorly active when extract was used independently of venom (pre-, co-, or posttreatment protocols). ∗∗∗Active in preincubation tests and when used independently of venom (pre-, co-, or posttreatment protocols).Table 4
List of medicinal plants with inhibitory potential against local effects induced byBitissnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Amaranthaceae
Pupalia lappacea
Herbal
B. arietans
SVH
—
[82]
Amaryllidaceae
Crinum jagus
Bulb
B. arietans
—
Myotoxicity∗
[167]
Anacardiaceae
Lannea acida
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Sclerocarya birrea
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Spondias
m
o
m
b
i
n
#
Cortex, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Annonaceae
Annona
s
e
n
e
g
a
l
e
n
s
i
s
#
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Apocynaceae
Strophanthus speciosus
Radix
B. arietans
SVH
—
[82]
Araliaceae
Polyscias fulva
Cortex
B. arietans
SVH
—
[82]
Bignoniaceae
Kigelia africana
Cortex
B. arietans
PLA2, SVH
—
[82]
Bixaceae
Cochlospermum tinctorium
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Capparaceae
Capparis tomentosa
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Colchicaceae
Gloriosa
s
u
p
e
r
b
a
#
Radix
B. arietans
SVH
—
[82]
Combretaceae
Combretum
m
o
l
l
e
#
Folium
B. arietans
PLA2, proteolytic, SVH
—
[82]
Guiera senegalensis
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Ebenaceae
Diospyros mespiliformis
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Euphorbiaceae
Alchornea laxiflora
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Fabaceae
Bauhinia thonningii
Cortex, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Bobgunnia madagascariensis(syn.Swartzia madagascariensis)
Folium, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Burkea africana
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Dichrostachys cinerea
Folium
B. arietans
PLA2, proteolytic, SVH
—
[82]
Entada africana
Radix
B. arietans
SVH
—
[82]
Parkia biglobosa
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Stylosanthes erecta
Folium
B. arietans
SVH
—
[82]
Tamarindus
i
n
d
i
c
a
#
Cortex, folium
B. arietans
PLA2, proteolytic, SVH
—
[82]
Hypericaceae
Psorospermum corymbiferum
Cortex, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Hypoxidaceae
Molineria capitulata(syn.Curculigo recurvata)
Folium
B. arietans
SVH
—
[82]
Lamiaceae
Rotheca myricoides(syn.Clerodendrum myricoides)
Cortex
B. arietans
SVH
—
[82]
Teucrium kraussii
Aerial parts, cortex
B. arietans
SVH
—
[82]
Volkameria glabra(syn.Clerodendrum glabrum)
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Lauraceae
Cassytha filiformis
Herbal
B. arietans
SVH
—
[82]
Loganiaceae
Strychnos decussata
Radix
B. arietans
Proteolytic
—
[82]
Strychnos innocua
Folium
B. arietans
Proteolytic, SVH
—
[82]
Malvaceae
Dombeya quinqueseta
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Grewia mollis
Cortex, folium, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Sterculia setigera
Cortex
B. arietans
PLA2, SVH
—
[82]
Waltheria indica
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Menispermaceae
Cissampelos mucronata
Herbal
B. arietans
Proteolytic, PLA2
—
[82]
Moraceae
Ficus platyphylla
Folium
B. arietans
PLA2, SVH
—
[82]
Olacaceae
Ximenia americana
Folium
B. arietans
PLA2, proteolytic, SVH
—
[82]
Phyllanthaceae
Flueggea virosa(syn. Securinega virosa)
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Primulaceae
Maesa
l
a
n
c
e
o
l
a
t
a
#
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Rhamnaceae
Ziziphus mucronata
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Ziziphus spina-christi
Cortex
B. arietans
PLA2, proteolytic, SVH
—
[82]
Rubiaceae
Crossopteryx febrifuga
Cortex
B. arietans
PLA2, SVH
—
[82]
Pentanisia prunelloides
Radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Pentas zanzibarica
Folium
B. arietans
PLA2
—
[82]
Rutaceae
Zanthoxylum capense
Radix
B. arietans
PLA2, proteolytic
—
[82]
Sapindaceae
Paullinia pinnata
Folium, radix
B. arietans
PLA2, proteolytic, SVH
—
[82]
Solanaceae
Schwenckia americana
Folium
B. arietans
SVH
—
[82]
Verbenaceae
Lantana trifolia
Cortex
B. arietans
PLA2, SVH
—
[82]
Vitaceae
Cissus populnea
Stem
B. arietans
SVH
—
[82]
PLA2 = snake venom phospholipase A2; SVH = snake venom hyaluronidase. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details).Table 5
List of medicinal plants with inhibitory potential against local effects induced byDaboia/Viperasnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Anacardiaceae
Anacardium
o
c
c
i
d
e
n
t
a
l
e
#
Bark
D. russelli
PLA2, proteolytic, SVH
Edema, hemorrhage, myotoxicity
[170]
Mangifera
i
n
d
i
c
a
#
Stem bark
D. russelii
LAAO, PLA2, SVH, proteolytic
Edema∗, hemorrhage∗, myotoxicity∗
[171]
Apocynaceae
Hemidesmus
i
n
d
i
c
u
s
#
Root
D. russelli
—
Hemorrhage
[172]
Tylophora
i
n
d
i
c
a
#
Leaf, root
D. russelli
PLA2
Hemorrhage
[85]
Aristolochiaceae
Aristolochia
b
r
a
c
t
e
o
l
a
t
a
#
Leaf, root
D. russelli
PLA2
Hemorrhage
[85]
Aristolochia
i
n
d
i
c
a
#
Root
D. russelii
LAAO, proteolytic
—
[173]
Asteraceae
Pluchea
i
n
d
i
c
a
#
Root
D. russelli
—
Hemorrhage
[172]
Euphorbiaceae
Acalypha
i
n
d
i
c
a
#
Leaf
D. r. russelli
—
Hemorrhage, necrosis
[174]
Fabaceae
Butea
m
o
n
o
s
p
e
r
m
a
#
Stem bark
D. russelii
SVH
Hemorrhage∗
[175]
Mimosa
p
u
d
i
c
a
#
Root
D. russelii
Proteolytic, SVH
—
[91]
Tamarindus
i
n
d
i
c
a
#
Seed
D. r. siamensis, D. russelii
LAAO, PLA2, SVH, proteolytic
Edema∗, hemorrhage∗, myotoxicity∗
[176, 177]
Lamiaceae
Leucas
a
s
p
e
r
a
#
Leaf, root
D. russelii
PLA2
Hemorrhage
[85]
Vitex
n
e
g
u
n
d
o
#
Root
D. russelii
—
Edema, hemorrhage
[178]
Loganiaceae
Strychnos
nux
-vomica
#
Seed
D. russelii
PLA2
Hemorrhage∗
[95]
Moraceae
Morus
a
l
b
a
#
Leaf
D. russelii
Proteolytic, SVH
Edema∗, hemorrhage∗, myotoxicity∗
[179]
Phyllanthaceae
Phyllanthus emblica(syn.Emblica officinalis)#
Root
D. russelii
—
Edema, hemorrhage
[178]
Piperaceae
Piper
l
o
n
g
u
m
#
Fruit
D. russelii
Hemorrhage
Edema, hemorrhage, myotoxicity, necrosis
[104]
Rubiaceae
Ophiorrhiza
m
u
n
g
o
s
#
Root
D. russelii
Hemorrhage
—
[180]
Salvadoraceae
Azima tetracantha
Leaf
D. russelii
SVH
—
[181]
Vitaceae
Vitis vinifera
Seed
D. russelii
Proteolytic, SVH
Edema∗, hemorrhage∗, myonecrosis∗
[182]
LAAO = L-amino acid oxidase; PLA2 = snake venom phospholipase A2; SVH = snake venom hyaluronidase. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details).Table 6
List of medicinal plants with inhibitory potential against local effects induced byLachesissnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Apocynaceae
Fernaldia pandurata(syn. Mandevilla velutina)
Root
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Asteraceae
Eclipta prostrata(syn.Eclipta alba)#
Aerial parts, root
L. muta
Myotoxicity, proteolytic, PLA2
Hemorrhage∗, myotoxicity
[130, 188]
Mikania glomerata
Root
L. muta
Proteolytic, PLA2
—
[188]
Erythroxylaceae
Erythroxylum ovalifolium
Stem
L. muta
Proteolytic, PLA2
Edema∗∗∗, hemorrhage∗∗∗
[189]
Erythroxylum subsessile
Stem
L. muta
Proteolytic, PLA2
Edema∗∗∗, hemorrhage∗∗∗
[189]
Euphorbiaceae
Jatropha elliptica
Root, stem
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Fabaceae
Pentaclethra
m
a
c
r
o
l
o
b
a
#
Bark
L. muta
—
Hemorrhage∗
[145]
Stryphnodendron adstringens(syn.Stryphnodendron barbatimam)
Root
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Melastomataceae
Miconia albicans
Stem
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Miconia fallax
Stem
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Miconia sellowiana
ND
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Tibouchina stenocarpa
Root
L. muta
Proteolytic, PLA2
Hemorrhage∗
[188]
Salicaceae
Casearia
s
y
l
v
e
s
t
r
i
s
#
Root
L. muta
Proteolytic
Hemorrhage∗
[188]
Sapotaceae
Manilkara subsericea
Leaf, stem
L. muta
Proteolytic, PLA2
Edema∗∗, hemorrhage∗∗
[100]
ND = information not described in the work; PLA2 = snake venom phospholipase A2. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details). ∗∗Active in preincubation tests but inactive or only poorly active when extract was used independently of venom (pre-, co-, or posttreatment protocols). ∗∗∗Active in preincubation tests and when used independently of venom (pre-, co-, or posttreatment protocols).Table 7
List of medicinal plants with inhibitory potential against local effects induced byCrotalussnakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Apocynaceae
Fernaldia pandurata(syn.Mandevilla velutina)
Leaf, stem, subterranean system
C. d. terrificus
PLA2
Edema∗, myotoxicity∗
[125]
Mandevilla illustris
Subterranean system
C. d. terrificus
PLA2
—
[190]
Asteraceae
Eclipta prostrata(syn.Eclipta alba)#
Aerial parts
C. d. terrificus
Myotoxicity
Myotoxicity∗
[11]
Bignoniaceae
Fridericia chica(syn.Arrabidaea chica)
Leaf
C. d. ruruima
—
Edema
[133]
Fabaceae
Pentaclethra
m
a
c
r
o
l
o
b
a
#
Bark
C. atrox
—
Hemorrhage∗
[145]
Schizolobium parahyba
Leaf
C. d. terrificus
PLA2
Edema∗
[146, 147]
Musaceae
M
u
s
a
×
p
a
r
a
d
i
s
i
a
c
a
#
Exudate
C. d. terrificus
PLA2
—
[155]
Rubiaceae
Randia
a
c
u
l
e
a
t
a
#
Fruit
C. simus
—
Myotoxicity
[78]
Sapindaceae
Sapindus saponaria
In vitro cultivated callus
C. d. terrificus
PLA2
—
[161]
PLA2 = snake venom phospholipase A2. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details).Table 8
List of medicinal plants with inhibitory potential against local effects induced by other snakes.
Plant name
Part used
Snake venom
Inhibited activities
Reference(s)
In vitro
In vivo
Amaryllidaceae
Crinum jagus
Bulb
Echis ocellatus
Hemorrhage
Myotoxicity∗∗
[167, 191]
Asteraceae
Artemisia absinthium
Aerial parts
Montivipera xanthina
—
Edema
[192]
Mikania laevigata
Leaf
Philodryas olfersii
Inflammation, myotoxicity
—
[193]
Fabaceae
Albizia
l
e
b
b
e
c
k
#
Seed
Echis carinatus
Proteolytic, SVH
Hemorrhage, myotoxicity
[194]
Mimosa
p
u
d
i
c
a
#
Root
Bungarus caeruleus, Echis carinatus
PLA2, proteolytic, SVH
Edema∗
[91, 92]
Parkia biglobosa
Stem bark
Echis ocellatus
Cytotoxicity against muscle cells, hemorrhage
—
[93]
Pentaclethra
m
a
c
r
o
l
o
b
a
#
Bark
Calloselasma rhodostoma
—
Hemorrhage∗
[145]
Senna auriculata(syn.Cassia auriculata)
Leaf
Echis carinatus
PLA2, proteolytic, SVH
Edema∗∗∗, hemorrhage∗∗∗, myotoxicity∗∗∗
[195]
Malvaceae
Hibiscus aethiopicus
Whole plant
Echis carinatus, Echis ocellatus
Cytotoxicity against muscle cells, hemorrhage
Edema∗∗∗, hemorrhage∗∗∗
[196, 197]
Salvadoraceae
Azima tetracantha
Leaf
Bungarus caeruleus
PLA2
—
[181]
Vitaceae
Vitis vinifera
Seed
Echis carinatus
Proteolytic, SVH
Edema∗, hemorrhage∗, myotoxicity∗
[198]
PLA2 = snake venom phospholipase A2; SVH = snake venom hyaluronidase. #Vegetal species with related folk use as antiophidic agents, as showed in Table 1. ∗Studies where inhibitory activity was assessed only by preincubation of venom with extract (see Section 4.1 for details). ∗∗Active in preincubation tests but inactive or only poorly active when extract was used independently of venom (pre-, co-, or posttreatment protocols). ∗∗∗Active in preincubation tests and when used independently of venom (pre-, co-, or posttreatment protocols).Along our survey were found 150 botanical families containing plants with reputation against snakebites, among which the most cited ones were the families Fabaceae, Asteraceae, Apocynaceae, Lamiaceae, Rubiaceae, Euphorbiaceae, Araceae, Malvaceae, and Acanthaceae (Figure1(a)). In a cross-cultural comparison of medicinal floras used against snakebites, Molander et al. [80] identified five countries with a high number of antiophidic plants and representing different cultures, geography, and floristic zones: Brazil, Nicaragua, Nepal, China, and South Africa. From these countries, some “hot” families were identified, which were Apocynaceae, Lamiaceae, Rubiaceae, and Zingiberaceae [80], similar to the present review, except for the Zingiberaceae family which was not so reported in our survey.Figure 1
“Hot families” with antiophidic potential. Main related botanical families in ethnopharmacological surveys as antiophidic (a) and main botanical families that were evaluated in antiophidic assay (inhibition of local tissue damage) and presented positive results (b).
(a)
(b)Medicinal plants with reputation against snakebites are found all over the world, especially in tropical or subtropical regions of Asia, Americas, and Africa (Figure2). This fact may be associated with richness of flora of these regions, as well as with relative need of complementary therapies to treat snakebites, considering geographical features that could limit the distribution and availability of the antivenoms in these areas.Figure 2
Distribution of medicinal plants used against snakebite around the world. World map highlighting the countries where antiophidic plants were related in ethnopharmacological surveys (a) and number of vegetal species per continent (b).
(a)
(b)As observed in Figure3(a), leaves and roots are the parts of plants most used in folk medicine. Regarding the mode of use, the most frequent one is the topical application of the vegetal products directly on the place of the bite (Figure 3(b)). This is interesting especially in snake venoms that cause serious local tissue damage, such asBothrops andDaboia species. Since these snakes produce intense local tissue damage, which has a very rapid onset, a topical treatment could be interesting for a rapid inhibitory action. On the other hand, interestingly, the use of some plant species is made by internal and external routes simultaneously, while for some other species the route of administration could be chosen among internal or external use. However, since in several cases this information is not clear, this differentiation was not considered in data tables. Regarding the mode of preparation, in general, paste and decoction were the most cited forms of use. However, for most of the plants enlisted, the information of mode of preparation was missing or confusing.Figure 3
Mode of utilization of antiophidic plants reported by folk medicine. Main plant parts used (a) and Venn diagram showing the number of species enlisted having external use, internal use, or both (b).
(a)
(b)It is important to emphasize that these plant species, in addition to their use as antiophidic agents, present a series of another popular uses (data not shown) in popular medicine, mainly anti-inflammatory activity. For example,Jatropha gossypiifolia (Euphorbiaceae) has antiophidic, anti-inflammatory, analgesic, antipyretic, healing, and antihemorrhagic uses, among others [81].


## 4. Antivenom Activities of Extracts of Medicinal Plants against Snake Venom Induced Local Tissue Damage

### 4.1. General Aspects

Until date, according to our database, only a few numbers (less than 20%) of the species with reputation against snakebites were tested in preclinical assays with different snake venoms, which shows that there is still a great road for the study of antiophidic plants. From these tested plants which have popular use documented in our database, more than a half (almost 60%) showed positive results, which shows that in fact ethnobotany could be a good tool for bioprospecting of plants with antiophidic activity. In addition, the fact that among the tested vegetal species very significative results were obtained strongly suggests the potentiality of these natural products as a future source for development of snake venom inhibitors.The plant families with most vegetal species showing positive results in antiophidic tests were Fabaceae, Euphorbiaceae, Apocynaceae, Lamiaceae, Asteraceae, Malvaceae, Melastomaceae, and Sapindaceae (Figure1(b)). Crossing the data of popular use (Figure 1(a)) and of positive activity (Figure 1(b)), we can highlight these families as “hot” ones, that is, families that might be preferred or prioritized in studies searching for antiophidic plants.Snakes from the genusNaja,Bothrops, andBitis were the most evaluated ones in these antiophidic assays. However, althoughNaja andBitis comprise a large fraction of the studies, virtually most of them are onlyin vitro studies, dealing with thein vitro enzymatic inhibition of classes of venom toxins relevant to local tissue damage, such as phospholipases A2 (PLA2s), hyaluronidases (SVHs), and proteases. More particularly, the great majority of these studies withNaja andBitis snakes are part of the work undertaken by Molander et al. [82], aiming to investigate whether plants used in traditional medicine systems would be active against necrosis-inducing enzymes of snake venoms, having tested a total of 226 extracts from 94 plants from the countries of Mali, Democratic Republic of Congo, and South Africa against PLA2, SVHs, and proteases fromBitis arietans andNaja nigricollis (see Tables 2 and 4). Studies evaluating the inhibitory action of medicinal plants against these enzymes are very relevant, since they are involved in several pathological mechanisms produced by snake venoms; however,in vivo preclinical assays or, even better, clinical assays are essential for giving even stronger evidences of the effectivity of the use of medicinal plants against snakebites. In this scenario, the study of anti-Bothrops plants is more advanced, since quantitatively a higher number ofin vivo scientific evidences are found in literature. Going the same way, studies with plants inhibiting local tissue damage ofDaboia/Vipera,Lachesis, andCrotalus snakes could be also highlighted. However, studies of antiophidic medicinal plants in humans are very scarce: only one clinical study was found in literature, evaluating the inhibitory properties of a polyherbal formulation against local effects from Chinese cobra bite (see Section 4.9).Hereafter, we describe the main plants with inhibitory potential against local tissue damage induced by snake venoms. It is important to emphasize that the focus of this review is plants against local tissue damage, mainly due to severity of these effects (which could cause permanent disabilities in victims) and the poor effectiveness of available antivenoms against them. So, studies with plants against systemic effects induced by snake were not considered; in addition some plants herein described possess inhibitory action upon systemic effects, although not stated here. For example, the vegetal speciesJatropha gossypiifolia (Euphorbiaceae), a medicinal plant studied very much by our research group, had showed significative inhibitory action upon hemostatic disorders induced byB. jararaca snake venom [96]. So, the antiophidic potential of this species (as well as some others) lies beyond the capacity of inhibit local tissue damage provoked byB. jararaca venom, although not described in this review.In addition, it is important to analyze critically some works dealing with antiophidic activity of plant extracts, since some of them have limitations that could reduce, at least partially, the potentiality of these species. The major limitation is that various studies, especially the early ones, make the evaluation of the plants using a preincubation approach, which consists in the previous inactivation of venom by preincubating it with different proportions of the tested extracts. Although scientifically valid and even recommended by WHO for assessing antiophidic antivenoms [97], this preincubation approach makes a scenario unlikely to be possible in the field, where the medicine would be delivered after the snakebite. In fact, a recent study evaluated the inhibitory action of the medicinal plantBellucia dichotoma (Melastomataceae) againstBothrops atrox snake venom using different protocols: preincubation, pretreatment, and posttreatment [98]. The authors observed that while the extract was greatly active when preincubated, this inhibitory activity was drastically reduced or even lost when the extract was injected independently of venom, simulating traditional use. The authors observed that the extract has great amounts of tannins, which are compounds known to precipitate proteins. So, it was concluded that the “pseudo-inhibition” observed after preincubation may be due to the presence of these compounds, suggesting that the preincubation protocol overestimates inhibitory potential of medicinal plants, and for this reason, this kind of approach must be analyzed with caution for estimation of inhibitory potential of medicinal plants [13, 98]. In this sense, many recent studies have been done using protocols of pre- and/or posttreatment, to ensure the potentiality of antiophidic plants, and for most of them, positive results have been found [96, 98–102]. For this reason, studies using preincubation protocol are marked in the tables, for a critical analysis.Also, it is interesting to note that several of the plants with inhibitory potential against snake venom local toxicities also present other relevant pharmacological activities. This is interesting since it is often discussed in the literature that several antiophidic plants did not neutralize snake venoms per se, but could have antiophidic use once they could relieve some of the symptoms of snake envenoming, especially the local effects. It is related that the presence of tranquilizing, antioxidant, immunostimulating, and/or anti-inflammatory activities in certain plants could be of great interest in the alleviation of snake envenoming symptoms [103, 104]. For example, some studies have shown that anti-inflammatory drugs could inhibit the edematogenic and other snake venom effects related to inflammation, such as necrosis and myotoxicity, induced byBothrops venoms [105, 106]. In fact, many medicinal plants with antiophidic activity also possess significant anti-inflammatory activityin vivo [83, 96, 107–110]. Following the same reasoning, some plants with antioxidant activity also possess significant antiophidic effects [95, 96, 104, 111]. In fact, some authors suggest that molecules with antioxidant and/or anti-inflammatory effects could be interesting along with antivenom therapy, helping to reduce the occurrence of secondary/long term complication due to snakebites [112].Bacterial infection secondary to snakebites is a common complication in envenomed victims [113, 114]. The main source of bacteria is the oral cavity of snakes, but the microbiota in the different layers of the victim’s skin or even microorganisms from victim’s clothes could also contribute [115, 116]. Abscess formation is a common complication found in patients bitten by Viperidae snakes, being a risk factor for amputation in these patients, and it may be associated with sepsis [113, 114, 117]. A large number of bacteria, including anaerobic species, aerobic gram-negative rods, and a small proportion of gram-positive cocci could be inoculated with snakebites and have been isolated from the abscesses of bitten patients [113, 114]. Microorganisms such asStaphylococcus,Pseudomonas,Salmonella,Escherichia,Providencia,Proteus,Enterococcus, andBacillus were already identified in oral cavity of certain snakes [116]. The use of antibiotics following snakebites is often recommended, usually therapeutically than prophylactically, mainly to avoid complications due to infections [114, 118]. In this context, medicinal plants presenting antimicrobial activities, especially against those microorganisms usually detected in snakebite victims’ abscesses, could be interesting [115].Medicinal plants having antimicrobial activities in association with some of the pharmacological properties discussed above (such as anti-inflammatory and antioxidant, e.g.) could be of great value to relieve especially local effects induced by snake venom. In another point of view, it is possible that several related plants in folk medicine as antiophidic agents do not act directly upon venom toxins but indirectly on its symptoms. Anyway, some studies have shown the potentiality of some vegetal species acting in two ways: directly, neutralizing venom toxins, or indirectly, by having some of the pharmacological activities mentioned above. For example,Jatropha gossypiifolia (Euphorbiaceae), a plant species studied very much in our research group, showed significant antiophidic properties, inhibiting biological and enzymatic activities fromBothrops venoms [96, 119], and presented anti-inflammatory, antioxidant, anticoagulant, and antimicrobial properties in preclinical assays [81]. So, plants which possess these biological activities determined in previous studies might be preferred or prioritized in studies searching for antiophidic plants.The mechanism by which medicinal plants neutralize the toxic venom constituents is still unknown, but many hypotheses have been proposed, such as protein precipitation, enzyme inactivation, proteolytic degradation, metal chelation, antioxidant action, and a combination of these mechanisms [15]. In this context, some improvements in this understanding have been achieved in the last years, through the use ofin silico methods (e.g., docking simulations) to analyze the interaction of compounds isolated from plants and certain classes of snake venom toxins such as PLA2 and SVMP [120–122].The use of medicinal plants may present several advantages, such as low cost, being easily available, being stable at room temperature, and possibility of neutralization of a wide range of venom components [15]. In addition, since medicinal plants are an extremely complex mixture, it is possible that there may be a synergistic action of different compounds in plant, acting in distinct targets, inhibiting a broad spectrum of venom toxins [12, 15]. According to literature, interestingly, there are some plants in which the crude extract is more active than the isolated constituents [15], which supports the hypothesis of the synergistic action of plant components.


### 4.2. Plants Inhibiting Naja Snakes

A summary of active plants againstNaja snakes local effects is presented in Table 2.Naja species are commonly called cobras. They typically occur in regions throughout Africa and Southern Asia. The outcomes of venom toxicity include nephro-, neuro-, and cardiotoxicity, respiratory and circulatory collapse, necrosis, hemorrhage, and edema [13]. A great number of the plants showed in this review were tested againstNaja species. However, it is important to mention that only a very small number of these plants were assessedin vivo, and so the scientific evidences of antiophidic activities of these species are based on enzymaticin vitro assays, especially against SVHs, a class of toxin particularly relevant in cobras. The study of Molander et al. [82] presented several medicinal plants identified as potent inhibitors ofN. nigricollis SVHs, PLA2, and proteases, which could indicate a potential rich source of inhibitors of necrosis induced by these venom, which must be evaluatedin vivo later [82]. The same group, in a more recent study [123], investigated the skin permeation,ex vivo inhibition of venom induced tissue destruction, and wound healing potential of African plants used against snakebite, which included the most potent inhibitors identified in the previous work [82]. A total of 30 plant species were tested againstNaja nigricollis andBitis arietans employingin vitro andex vivo models [123]. However, although plant extracts have showed potential in inhibiting snake venom enzymes, this study showed no effect against cell death and tissue damage.


### 4.3. Plants Inhibiting Bothrops Snakes

A summary of active plants againstBothrops snakes local effects is presented in Table 3. More than 90% of the snakebites reported every year in Latin America are caused byBothrops species [8]. Envenomation byBothrops snakes is characterized by a prominent and complex series of local pathological alterations, which appear rapidly after the bite in the anatomical site where venom is inoculated [168]. In a number ofBothrops bite cases, lack of neutralization of local effects results in permanent sequelae, with significative tissue loss [8]. So, the use of a therapeutic approach with high inhibitory potential and easy access and disponibility to victims, which could neutralize rapidly the onset of these local manifestations, is interesting. Most of the inhibitory studies withBothrops snakes were performed in Brazil, which could be associated with richness of Brazilian flora as well as the epidemiological aspects of this country. The work performed by De Moura et al. [33] could be highlighted, where these authors performed an ethnopharmacological-guided screening of plants with reputation against snakebite in Santarém, Western Pará, Brazil. Twelve species were evaluated againstBothrops jararaca snake venom induced hemorrhage and some of them presented very significative results, showing, thus, the relevance of traditional knowledge in the survey of antiophidic plants [33].


### 4.4. Plants Inhibiting Bitis Snakes

A summary of active plants againstBitis snakes local effects is presented in Table 4. Snakes belonging to the genusBitis are implicated in many accidents with humans in Africa. The envenomation byBitis often results in severe local damage, hypotension, coagulopathy, thrombocytopenia, and spontaneous local bleeding and, in the absence of antivenom therapy, the accident can be fatal.Bitis arietans is one of the three species of snakes of medical importance in Africa and its venom is considered the most toxic venom of the viper group [169]. Regarding the plants with inhibitory action uponBitis snakes, only onein vivo study of antiophidic activity was found until date. Although many works have been showing the potential of medicinal plants against several snake venoms, only three works were identified evaluating the action of plants againstBitis, from which two are the same screening studies of plants againstNaja snake venom discussed before (Section 4.2) [82, 123].


### 4.5. Plants Inhibiting Daboia/Vipera Snakes

A summary of active plants againstDaboia/Vipera snakes local effects is presented in Table 5. TheDaboia genus is represented by a single species, namedDaboia russelii, also popularly known as Russell’s viper. This species is widespread in many parts of Asia and is responsible for large morbimortality due to snakebites in this continent [183, 184]. Russell’s viper was formerly classified inVipera genus and is therefore better known asVipera russelii, since the new accepted nomenclature(Daboia russelii) is not yet universally followed [184]. For this reason, to avoid confounding, we use the termDaboia/Vipera in some occasions.In humans, Russell’s viper bite causes severe local tissue damage; more frequently the necrosis results in an irreversible loss of tissue and requires amputation of the affected limb [182, 183, 185]. As observed withBothrops snakes, several studies have showed the inhibitory potential of medicinal plants against local effects of Russell’s viper venom, including several preclinicalin vivo studies.


### 4.6. Plants Inhibiting Lachesis Snakes

A summary of active plants againstLachesis snakes local effects is presented in Table 6.Lachesis muta is the longest venomous snake in the Americas and is distributed in the equatorial forests east of the Andes, ranging from eastern Ecuador, Colombia, Peru, northern Bolivia, and eastern and northern Venezuela, to Guyana, French Guyana, Surinam, and northern Brazil [100, 186].L. muta snakebites are mainly characterized by systemic (generalized bleeding, coagulopathy, renal failure, and shock) and local effects (pain, hemorrhage, edema, and necrosis). In South America,Bothrops species has a higher incidence of accidents thanL. muta, but, on the other hand,Lachesis bites led to more severe symptoms and have lethality indexes significantly higher thanBothrops [100, 186, 187]. Thus, the study of medicinal plants against these snakes, too, is of very much relevance. However, only a few studies were detected with plants againstLachesis snakes.


### 4.7. Plants Inhibiting Crotalus Snakes

A summary of active plants againstCrotalus snakes local effects is presented in Table 7. Snakes fromCrotalus durissus complex, popularly known as rattlesnakes, are dispersed northward into North America and southward into South America. Species of theCrotalus durissus complex pose a serious medical problem in many parts of the America [199]. Crotalic venom is considered highly toxic and more lethal in comparison with that of the genusBothrops, having three main actions: neurotoxic, myotoxic, and coagulant [200, 201]. The crotalic accident is characterized by local and systemic manifestations, but while the local alterations are only discrete, the systemic manifestations are severe, leading to high chances of death [201]. Probably due to this low local effect in envenomed victims, the inhibition of these effects by plants is, until now, little investigated, especially when compared to other species with characteristic severe local effects.


### 4.8. Plants Inhibiting Other Snakes

Besides the snakes discussed above, some other studies are found with plants inhibiting other snake species, such as those fromEchis andBungarus genus. For other snakes species such asCalloselasma rhodostoma,Philodryas olfersii, andMontivipera xanthina, only isolated studies with a single plant, in each one, were found. These plants are summarized in Table 8. Many reasons may be stated for this lack of studies, such as low level of local effects, incidence restricted to a small region of the world, and usual low efficacy of plant extracts due to possible extremely high toxicity. However, it is important to highlight that the lack of studies does not mean a lower medical relevance of these species. For example, the saw-scaled viper(Echis carinatus) and the common Indian krait(Bungarus caeruleus), along with spectacled cobra(Naja naja) and Russell’s viper(Daboia russelii), are included among the referred “Big Four” venomous snakes of India, being responsible for the majority of morbid complications, characterized by persistent and progressive tissue necrosis even after treatment with antivenom [195, 202]. Therefore, future studies with plants aiming at the inhibition of the local effects induced by these snakes are encouraged.


### 4.9. Studies in Humans

Along our antiophidic plants database, only one clinical study was found in literature, evaluating the inhibitory properties of a polyherbal formulation, externally applied, against soft-tissue necrosis afterNaja atra (Chinese cobra) bite [203]. This polyherbal formulation, known in China as Jidesheng antivenom, is composed of the following ingredients: Ganchan(Succys Bufo), Dijincao(Herba Euphorbiae Humifusae), Chonglou(Rhizoma Paridis Chonglou), and Wugong(Scolopendra). This was a retrospective study performed with 126 patients with skin and soft-tissue necrosis due cobra bite, with the control group being treated externally with 40% glyceride magnesium sulfate (n=52) and the treatment group performed by application of Jidesheng antivenom externally (n=74). The authors observed statistically significant differences in maximum local necrotic area of skin and soft tissues, healing time, and skin-grafting rate between the control and treatment groups (P<0.05), thus indicating that external application of Jidesheng antivenom may help to promote wound healing and reduce the skin-grafting rate in cases of skin and soft-tissue necrosis due to Chinese cobra bite [203]. Considering the composition of the Jidesheng antivenom, the authors discuss that each ingredient in this product may exert antipyretic, antidotal, antiphlogistic, and analgesic effects, according to previous results with each ingredient isolated, which could contribute to the inhibitory effect observed by the formulation [203]. The result obtained in this clinical study is very promising, since it shows that a plant-derived product showed significant results in humans, thus pointing to the potentiality of this kind of product in treatment of snake venom induced local effects. However, only one study is insufficient to ensure the potentiality of medicinal plants against snakebites, with performing more clinical studies, preferentially controlled and randomized ones, to bring more evidences of the viability of the approach for future safe and effective use in humans being necessary. So, more clinical studies, especially ones with those plants highlighted in this review and those presenting good preclinicalin vivo evidences of antiophidic efficacy, are highly encouraged.


## 4.1. General Aspects

Until date, according to our database, only a few numbers (less than 20%) of the species with reputation against snakebites were tested in preclinical assays with different snake venoms, which shows that there is still a great road for the study of antiophidic plants. From these tested plants which have popular use documented in our database, more than a half (almost 60%) showed positive results, which shows that in fact ethnobotany could be a good tool for bioprospecting of plants with antiophidic activity. In addition, the fact that among the tested vegetal species very significative results were obtained strongly suggests the potentiality of these natural products as a future source for development of snake venom inhibitors.The plant families with most vegetal species showing positive results in antiophidic tests were Fabaceae, Euphorbiaceae, Apocynaceae, Lamiaceae, Asteraceae, Malvaceae, Melastomaceae, and Sapindaceae (Figure1(b)). Crossing the data of popular use (Figure 1(a)) and of positive activity (Figure 1(b)), we can highlight these families as “hot” ones, that is, families that might be preferred or prioritized in studies searching for antiophidic plants.Snakes from the genusNaja,Bothrops, andBitis were the most evaluated ones in these antiophidic assays. However, althoughNaja andBitis comprise a large fraction of the studies, virtually most of them are onlyin vitro studies, dealing with thein vitro enzymatic inhibition of classes of venom toxins relevant to local tissue damage, such as phospholipases A2 (PLA2s), hyaluronidases (SVHs), and proteases. More particularly, the great majority of these studies withNaja andBitis snakes are part of the work undertaken by Molander et al. [82], aiming to investigate whether plants used in traditional medicine systems would be active against necrosis-inducing enzymes of snake venoms, having tested a total of 226 extracts from 94 plants from the countries of Mali, Democratic Republic of Congo, and South Africa against PLA2, SVHs, and proteases fromBitis arietans andNaja nigricollis (see Tables 2 and 4). Studies evaluating the inhibitory action of medicinal plants against these enzymes are very relevant, since they are involved in several pathological mechanisms produced by snake venoms; however,in vivo preclinical assays or, even better, clinical assays are essential for giving even stronger evidences of the effectivity of the use of medicinal plants against snakebites. In this scenario, the study of anti-Bothrops plants is more advanced, since quantitatively a higher number ofin vivo scientific evidences are found in literature. Going the same way, studies with plants inhibiting local tissue damage ofDaboia/Vipera,Lachesis, andCrotalus snakes could be also highlighted. However, studies of antiophidic medicinal plants in humans are very scarce: only one clinical study was found in literature, evaluating the inhibitory properties of a polyherbal formulation against local effects from Chinese cobra bite (see Section 4.9).Hereafter, we describe the main plants with inhibitory potential against local tissue damage induced by snake venoms. It is important to emphasize that the focus of this review is plants against local tissue damage, mainly due to severity of these effects (which could cause permanent disabilities in victims) and the poor effectiveness of available antivenoms against them. So, studies with plants against systemic effects induced by snake were not considered; in addition some plants herein described possess inhibitory action upon systemic effects, although not stated here. For example, the vegetal speciesJatropha gossypiifolia (Euphorbiaceae), a medicinal plant studied very much by our research group, had showed significative inhibitory action upon hemostatic disorders induced byB. jararaca snake venom [96]. So, the antiophidic potential of this species (as well as some others) lies beyond the capacity of inhibit local tissue damage provoked byB. jararaca venom, although not described in this review.In addition, it is important to analyze critically some works dealing with antiophidic activity of plant extracts, since some of them have limitations that could reduce, at least partially, the potentiality of these species. The major limitation is that various studies, especially the early ones, make the evaluation of the plants using a preincubation approach, which consists in the previous inactivation of venom by preincubating it with different proportions of the tested extracts. Although scientifically valid and even recommended by WHO for assessing antiophidic antivenoms [97], this preincubation approach makes a scenario unlikely to be possible in the field, where the medicine would be delivered after the snakebite. In fact, a recent study evaluated the inhibitory action of the medicinal plantBellucia dichotoma (Melastomataceae) againstBothrops atrox snake venom using different protocols: preincubation, pretreatment, and posttreatment [98]. The authors observed that while the extract was greatly active when preincubated, this inhibitory activity was drastically reduced or even lost when the extract was injected independently of venom, simulating traditional use. The authors observed that the extract has great amounts of tannins, which are compounds known to precipitate proteins. So, it was concluded that the “pseudo-inhibition” observed after preincubation may be due to the presence of these compounds, suggesting that the preincubation protocol overestimates inhibitory potential of medicinal plants, and for this reason, this kind of approach must be analyzed with caution for estimation of inhibitory potential of medicinal plants [13, 98]. In this sense, many recent studies have been done using protocols of pre- and/or posttreatment, to ensure the potentiality of antiophidic plants, and for most of them, positive results have been found [96, 98–102]. For this reason, studies using preincubation protocol are marked in the tables, for a critical analysis.Also, it is interesting to note that several of the plants with inhibitory potential against snake venom local toxicities also present other relevant pharmacological activities. This is interesting since it is often discussed in the literature that several antiophidic plants did not neutralize snake venoms per se, but could have antiophidic use once they could relieve some of the symptoms of snake envenoming, especially the local effects. It is related that the presence of tranquilizing, antioxidant, immunostimulating, and/or anti-inflammatory activities in certain plants could be of great interest in the alleviation of snake envenoming symptoms [103, 104]. For example, some studies have shown that anti-inflammatory drugs could inhibit the edematogenic and other snake venom effects related to inflammation, such as necrosis and myotoxicity, induced byBothrops venoms [105, 106]. In fact, many medicinal plants with antiophidic activity also possess significant anti-inflammatory activityin vivo [83, 96, 107–110]. Following the same reasoning, some plants with antioxidant activity also possess significant antiophidic effects [95, 96, 104, 111]. In fact, some authors suggest that molecules with antioxidant and/or anti-inflammatory effects could be interesting along with antivenom therapy, helping to reduce the occurrence of secondary/long term complication due to snakebites [112].Bacterial infection secondary to snakebites is a common complication in envenomed victims [113, 114]. The main source of bacteria is the oral cavity of snakes, but the microbiota in the different layers of the victim’s skin or even microorganisms from victim’s clothes could also contribute [115, 116]. Abscess formation is a common complication found in patients bitten by Viperidae snakes, being a risk factor for amputation in these patients, and it may be associated with sepsis [113, 114, 117]. A large number of bacteria, including anaerobic species, aerobic gram-negative rods, and a small proportion of gram-positive cocci could be inoculated with snakebites and have been isolated from the abscesses of bitten patients [113, 114]. Microorganisms such asStaphylococcus,Pseudomonas,Salmonella,Escherichia,Providencia,Proteus,Enterococcus, andBacillus were already identified in oral cavity of certain snakes [116]. The use of antibiotics following snakebites is often recommended, usually therapeutically than prophylactically, mainly to avoid complications due to infections [114, 118]. In this context, medicinal plants presenting antimicrobial activities, especially against those microorganisms usually detected in snakebite victims’ abscesses, could be interesting [115].Medicinal plants having antimicrobial activities in association with some of the pharmacological properties discussed above (such as anti-inflammatory and antioxidant, e.g.) could be of great value to relieve especially local effects induced by snake venom. In another point of view, it is possible that several related plants in folk medicine as antiophidic agents do not act directly upon venom toxins but indirectly on its symptoms. Anyway, some studies have shown the potentiality of some vegetal species acting in two ways: directly, neutralizing venom toxins, or indirectly, by having some of the pharmacological activities mentioned above. For example,Jatropha gossypiifolia (Euphorbiaceae), a plant species studied very much in our research group, showed significant antiophidic properties, inhibiting biological and enzymatic activities fromBothrops venoms [96, 119], and presented anti-inflammatory, antioxidant, anticoagulant, and antimicrobial properties in preclinical assays [81]. So, plants which possess these biological activities determined in previous studies might be preferred or prioritized in studies searching for antiophidic plants.The mechanism by which medicinal plants neutralize the toxic venom constituents is still unknown, but many hypotheses have been proposed, such as protein precipitation, enzyme inactivation, proteolytic degradation, metal chelation, antioxidant action, and a combination of these mechanisms [15]. In this context, some improvements in this understanding have been achieved in the last years, through the use ofin silico methods (e.g., docking simulations) to analyze the interaction of compounds isolated from plants and certain classes of snake venom toxins such as PLA2 and SVMP [120–122].The use of medicinal plants may present several advantages, such as low cost, being easily available, being stable at room temperature, and possibility of neutralization of a wide range of venom components [15]. In addition, since medicinal plants are an extremely complex mixture, it is possible that there may be a synergistic action of different compounds in plant, acting in distinct targets, inhibiting a broad spectrum of venom toxins [12, 15]. According to literature, interestingly, there are some plants in which the crude extract is more active than the isolated constituents [15], which supports the hypothesis of the synergistic action of plant components.


## 4.2. Plants Inhibiting Naja Snakes

A summary of active plants againstNaja snakes local effects is presented in Table 2.Naja species are commonly called cobras. They typically occur in regions throughout Africa and Southern Asia. The outcomes of venom toxicity include nephro-, neuro-, and cardiotoxicity, respiratory and circulatory collapse, necrosis, hemorrhage, and edema [13]. A great number of the plants showed in this review were tested againstNaja species. However, it is important to mention that only a very small number of these plants were assessedin vivo, and so the scientific evidences of antiophidic activities of these species are based on enzymaticin vitro assays, especially against SVHs, a class of toxin particularly relevant in cobras. The study of Molander et al. [82] presented several medicinal plants identified as potent inhibitors ofN. nigricollis SVHs, PLA2, and proteases, which could indicate a potential rich source of inhibitors of necrosis induced by these venom, which must be evaluatedin vivo later [82]. The same group, in a more recent study [123], investigated the skin permeation,ex vivo inhibition of venom induced tissue destruction, and wound healing potential of African plants used against snakebite, which included the most potent inhibitors identified in the previous work [82]. A total of 30 plant species were tested againstNaja nigricollis andBitis arietans employingin vitro andex vivo models [123]. However, although plant extracts have showed potential in inhibiting snake venom enzymes, this study showed no effect against cell death and tissue damage.


## 4.3. Plants Inhibiting Bothrops Snakes

A summary of active plants againstBothrops snakes local effects is presented in Table 3. More than 90% of the snakebites reported every year in Latin America are caused byBothrops species [8]. Envenomation byBothrops snakes is characterized by a prominent and complex series of local pathological alterations, which appear rapidly after the bite in the anatomical site where venom is inoculated [168]. In a number ofBothrops bite cases, lack of neutralization of local effects results in permanent sequelae, with significative tissue loss [8]. So, the use of a therapeutic approach with high inhibitory potential and easy access and disponibility to victims, which could neutralize rapidly the onset of these local manifestations, is interesting. Most of the inhibitory studies withBothrops snakes were performed in Brazil, which could be associated with richness of Brazilian flora as well as the epidemiological aspects of this country. The work performed by De Moura et al. [33] could be highlighted, where these authors performed an ethnopharmacological-guided screening of plants with reputation against snakebite in Santarém, Western Pará, Brazil. Twelve species were evaluated againstBothrops jararaca snake venom induced hemorrhage and some of them presented very significative results, showing, thus, the relevance of traditional knowledge in the survey of antiophidic plants [33].


## 4.4. Plants Inhibiting Bitis Snakes

A summary of active plants againstBitis snakes local effects is presented in Table 4. Snakes belonging to the genusBitis are implicated in many accidents with humans in Africa. The envenomation byBitis often results in severe local damage, hypotension, coagulopathy, thrombocytopenia, and spontaneous local bleeding and, in the absence of antivenom therapy, the accident can be fatal.Bitis arietans is one of the three species of snakes of medical importance in Africa and its venom is considered the most toxic venom of the viper group [169]. Regarding the plants with inhibitory action uponBitis snakes, only onein vivo study of antiophidic activity was found until date. Although many works have been showing the potential of medicinal plants against several snake venoms, only three works were identified evaluating the action of plants againstBitis, from which two are the same screening studies of plants againstNaja snake venom discussed before (Section 4.2) [82, 123].


## 4.5. Plants Inhibiting Daboia/Vipera Snakes

A summary of active plants againstDaboia/Vipera snakes local effects is presented in Table 5. TheDaboia genus is represented by a single species, namedDaboia russelii, also popularly known as Russell’s viper. This species is widespread in many parts of Asia and is responsible for large morbimortality due to snakebites in this continent [183, 184]. Russell’s viper was formerly classified inVipera genus and is therefore better known asVipera russelii, since the new accepted nomenclature(Daboia russelii) is not yet universally followed [184]. For this reason, to avoid confounding, we use the termDaboia/Vipera in some occasions.In humans, Russell’s viper bite causes severe local tissue damage; more frequently the necrosis results in an irreversible loss of tissue and requires amputation of the affected limb [182, 183, 185]. As observed withBothrops snakes, several studies have showed the inhibitory potential of medicinal plants against local effects of Russell’s viper venom, including several preclinicalin vivo studies.


## 4.6. Plants Inhibiting Lachesis Snakes

A summary of active plants againstLachesis snakes local effects is presented in Table 6.Lachesis muta is the longest venomous snake in the Americas and is distributed in the equatorial forests east of the Andes, ranging from eastern Ecuador, Colombia, Peru, northern Bolivia, and eastern and northern Venezuela, to Guyana, French Guyana, Surinam, and northern Brazil [100, 186].L. muta snakebites are mainly characterized by systemic (generalized bleeding, coagulopathy, renal failure, and shock) and local effects (pain, hemorrhage, edema, and necrosis). In South America,Bothrops species has a higher incidence of accidents thanL. muta, but, on the other hand,Lachesis bites led to more severe symptoms and have lethality indexes significantly higher thanBothrops [100, 186, 187]. Thus, the study of medicinal plants against these snakes, too, is of very much relevance. However, only a few studies were detected with plants againstLachesis snakes.


## 4.7. Plants Inhibiting Crotalus Snakes

A summary of active plants againstCrotalus snakes local effects is presented in Table 7. Snakes fromCrotalus durissus complex, popularly known as rattlesnakes, are dispersed northward into North America and southward into South America. Species of theCrotalus durissus complex pose a serious medical problem in many parts of the America [199]. Crotalic venom is considered highly toxic and more lethal in comparison with that of the genusBothrops, having three main actions: neurotoxic, myotoxic, and coagulant [200, 201]. The crotalic accident is characterized by local and systemic manifestations, but while the local alterations are only discrete, the systemic manifestations are severe, leading to high chances of death [201]. Probably due to this low local effect in envenomed victims, the inhibition of these effects by plants is, until now, little investigated, especially when compared to other species with characteristic severe local effects.


## 4.8. Plants Inhibiting Other Snakes

Besides the snakes discussed above, some other studies are found with plants inhibiting other snake species, such as those fromEchis andBungarus genus. For other snakes species such asCalloselasma rhodostoma,Philodryas olfersii, andMontivipera xanthina, only isolated studies with a single plant, in each one, were found. These plants are summarized in Table 8. Many reasons may be stated for this lack of studies, such as low level of local effects, incidence restricted to a small region of the world, and usual low efficacy of plant extracts due to possible extremely high toxicity. However, it is important to highlight that the lack of studies does not mean a lower medical relevance of these species. For example, the saw-scaled viper(Echis carinatus) and the common Indian krait(Bungarus caeruleus), along with spectacled cobra(Naja naja) and Russell’s viper(Daboia russelii), are included among the referred “Big Four” venomous snakes of India, being responsible for the majority of morbid complications, characterized by persistent and progressive tissue necrosis even after treatment with antivenom [195, 202]. Therefore, future studies with plants aiming at the inhibition of the local effects induced by these snakes are encouraged.


## 4.9. Studies in Humans

Along our antiophidic plants database, only one clinical study was found in literature, evaluating the inhibitory properties of a polyherbal formulation, externally applied, against soft-tissue necrosis afterNaja atra (Chinese cobra) bite [203]. This polyherbal formulation, known in China as Jidesheng antivenom, is composed of the following ingredients: Ganchan(Succys Bufo), Dijincao(Herba Euphorbiae Humifusae), Chonglou(Rhizoma Paridis Chonglou), and Wugong(Scolopendra). This was a retrospective study performed with 126 patients with skin and soft-tissue necrosis due cobra bite, with the control group being treated externally with 40% glyceride magnesium sulfate (n=52) and the treatment group performed by application of Jidesheng antivenom externally (n=74). The authors observed statistically significant differences in maximum local necrotic area of skin and soft tissues, healing time, and skin-grafting rate between the control and treatment groups (P<0.05), thus indicating that external application of Jidesheng antivenom may help to promote wound healing and reduce the skin-grafting rate in cases of skin and soft-tissue necrosis due to Chinese cobra bite [203]. Considering the composition of the Jidesheng antivenom, the authors discuss that each ingredient in this product may exert antipyretic, antidotal, antiphlogistic, and analgesic effects, according to previous results with each ingredient isolated, which could contribute to the inhibitory effect observed by the formulation [203]. The result obtained in this clinical study is very promising, since it shows that a plant-derived product showed significant results in humans, thus pointing to the potentiality of this kind of product in treatment of snake venom induced local effects. However, only one study is insufficient to ensure the potentiality of medicinal plants against snakebites, with performing more clinical studies, preferentially controlled and randomized ones, to bring more evidences of the viability of the approach for future safe and effective use in humans being necessary. So, more clinical studies, especially ones with those plants highlighted in this review and those presenting good preclinicalin vivo evidences of antiophidic efficacy, are highly encouraged.


## 5. Concluding Remarks

The popular use of vegetal species does not necessarily imply efficacy, but it gives a selected list of medicinal plants that can be primarily studied in pharmacologic assays for possible antiophidic effects, directing future studies in this area. In fact, a great number of these species that have been evaluated against local tissue damage induced by several snake species showed inhibitory potential against hyaluronidase, phospholipase, proteolytic, hemorrhagic, myotoxic, and edematogenic activities, among others. Therefore, considering the limitations of conventional antivenom serotherapy, especially its poor efficacy against local effects, the treatment with medicinal plants may provide a potential adjuvant alternative to treat snakebites, being used to complement the activity and effectiveness of available snake venom therapy. The main potential advantages of antiophidic plants are their low cost, easy access, stability at room temperature, and ability to neutralize a broad spectrum of toxins, including the local tissue damage.Interestingly, some studies have showed that the crude extracts are more powerful than the individual herbal compounds, which could, at a certain extent, justify the development of herbal products containing these plants instead of medicines containing isolated compounds, which in turn could be more rapidly available in market, after proof of safety, effectiveness, and quality of these products. However, despite the existence of many plants with great potential, no natural antiophidic product is available in market, which points to question of the need for further studies. Only a few numbers of patents regarding herbal products against snakebites were found in literature. Some patents regarding the use of Chinese medicinal plants against snake and bug bites were found. In our research group, two patents were deposited concerning the processes of obtaining extracts, fraction, isolated compounds, and pharmaceutical compositions of some plants studied by our group applied in the treatment of accidents with venomous animals (BR 10 2013 034046 4 A2 and BR 10 2012 026958 9 A2). Thus, the number of patents with antiophidic herbal products is still relatively small. For this reason, we encourage pharmacologists and toxinologists around the world to intensify studies with antiophidic plants, especially prioritizing those with the greatest number of indications in traditional medicine and emphasizing clinical studies with the most active plants in preclinical studies, given that the low number of human studies is one of the major obstacles for the future application of herbal products with antiophidic potential. No less important, toxicological studies are also extremely necessary to ensure the safety of these products.In conclusion, the data presented in this review provides an updated scenario for and insights into future research aiming at validation of medicinal plants as antiophidic agents and, based on scientific evidences, strengthens the potentiality of medicinal plants and ethnopharmacological knowledge as a tool for design of potent inhibitors and/or herbal medicines against venom toxins.



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