After Linnaeus, the British gardener Philip Miller (1768) recognised 10 domesticated species, five of them new species described by him. The descriptions of all these species were almost certainly made from living plants cultivated in the Chelsea Physic Garden in London.

The German botanist, Johann Heinrich Dierbach (1829), proposed a particularly confusing infraspecific classification. He accepted a single species and validated the pre-Linnaean name “Capsicum indicum” of L’Obel (1576), as C. indicum Dierb. He recognised infraspecific unranked taxa designated by numbers (1. macrocarpon, 2. pachycarb[p]on, 3. cerasocarpon, 4. elaeocarpon, 5. microcarpon), based on fruit shape; none of these numbered names had a direct reference to the author or a specimen. Within each numbered category, the different taxa were designated with a letter (a to e; e.g. C. indicum Lobel. 1. macrocarpon b. longum) and, in addition, before the designation of letters, the taxa were grouped according to the pericarp colour (e.g. pericarpiis rubris, pericarpiis flavis). Heiser and Pickersgill (1975) attempted to interpret Dierbach’s classification system, concluding that it “… should be ignored for nomenclatural purposes”. However, Dierbach based the lettered infraspecific taxa on the names of the chili peppers known at that time, with the result that many of them are now synonyms (especially under C. annuum) or as doubtful names (see Doubtful names).

The first illustrated monograph of Capsicum was published by the German botanist, Karl Anton Fingerhuth (1832), in which he included 32 species. He recognised the taxa already described by previous authors and described only three new species (C. cumanense Fingerh., C. strictum Fingerh. and C. ceratocarpum Fingerh.). He also recognised 29 varieties, most of which were newly described. Although he did not propose a formal infrageneric classification, he grouped the species into two groups (A and B) depending on the position of the fruits (A. Fruits erect; B. Fruits pendent). Within each group, he further separated the species into two subgroups (a. Fruits oblong; b. Fruits subglobose). The distinctions of his many varieties were based on the broad range of different fruit shapes. None of the taxa proposed by Fingerhuth is an accepted name in this treatment (see synonymy under the domesticated species).

In his monumental treatment of Solanaceae, Michel-Felix Dunal (1852) recognised 61 Capsicum species (16 newly described by him), together with 51 varieties (16 new), 11 species requiring further study and three doubtful names. From these, 44 species and 40 varieties referred to the domesticated species. Dunal did not establish an infrageneric classification, but grouped the species in a similar way as had Fingerhuth (1832), first by the position of the fruits (erect or pendent) and then by shape (oblong or globose-ovate).

In extreme contrast, other authors tended to reduce the number of accepted species to two (Irish 1898) or only one (Bailey 1923; Erwin 1929; Shinners 1956; and others), with very complex infraspecific hierarchical systems, based on the extreme variation in fruit characters of the domesticated taxa and their cultivars. Terpó (1966) summarised some of these elaborate systems, while Eshbaugh (1980) expressed the difficulties inherent in developing a useful and rational system that satisfied everyone. Capsicum annuum and C. frutescens and, to a lesser extent for C. chinense, have been the taxa most debated by taxonomists and horticulturists in classifications below the species level. From an evolutionary perspective, the relationships between these three species have been the most debated (Pickersgill 1988), whereas there was a general agreement that C. pubescens and C. baccatum var. pendulum were two well-defined domesticated taxa (Eshbaugh 1980; Pickersgill 1988).

As a crop genus, Capsicum has inspired researchers to follow a number of in-depth approaches since the mid-1900s, such as classical and molecular cytogenetic analyses, crossing experiments, biochemical and protein electrophoretic studies, molecular characterisation through genotypic markers (restriction fragment length polymorphism, RFLP, amplified fragment length polymorphism, AFLP, random amplified polymorphic DNA, RAPD, microsatellite or simple sequence repeat, SSR, random amplified microsatellite polymorphism, RAMPO and direct amplification of minisatellite DNA, DAMDPCR), phytogeographic and phylogeographic analyses, chloroplast and nuclear DNA and whole-genome sequencing studies (see Moscone et al. 2007; Aguilar-Meléndez et al. 2009; Carvalho et al. 2014; Carrizo García et al. 2016; Raveendar et al. 2017; Scaldaferro et al. 2018; Tripodi et al. 2019; and references in each). This evidence has contributed to a better understanding of the complex taxonomy of the domesticated (and wild) species, as well as to determine the origin of the crop species. Most current taxonomic works and breeding programmes recognise five domesticated species of Capsicum (Pickersgill 2016).

Taxonomic work on wild Capsicum species began in the 19^th century. Initially, some authors (Cavanilles 1802; Willdenow 1809; Dunal 1816; Kunth 1818; Roemer and Schultes 1819; and others) described isolated species. The Capsicum treatment by Sendtner (1846) in “Flora Brasiliensis” is the first significant work from an extended geographical area; he accepted seven wild species and five infraspecific taxa in addition to several cultivated species. Later, Dunal (1852) made an important contribution with the addition of seven other novel wild species and three varieties.

During the early 20^th century, sporadic descriptions of new Capsicum taxa continued (Chodat 1902; Chodat and Hassler 1903; Witasek 1910; Bitter 1919, 1921, 1922, 1924). Later, a new generation of taxonomists interested in this genus, led by Paul Smith and Charles Heiser, Jr. (in the United States) and Armando T. Hunziker (in Argentina), made great advances in the understanding of the generic limits of Capsicum (see above). These authors not only provided new names, but also new evidence to establish the relationships between species. Hunziker, especially, contributed greatly to a comprehensive understanding of the taxonomy of the genus (Hunziker 1950, 1956, 1961, 1969b, 1971, 1998, 2001). In his book “Genera Solanacearum”, Hunziker (2001) proposed Capsicum as a natural group with “ca. 20 species and a few varieties”.

In the last two decades, field explorations across South America, mainly in the central Andean countries and Brazil, have enabled us to gain a better understanding of the genus as a whole. Thirteen new wild species have been described and partial keys for the identification of the species for particular areas have been provided (Barboza and Bianchetti 2005; Nee et al. 2006; Barboza et al. 2011, 2019; Barboza et al. 2020a, Barboza et al. 2020b). We recognise here 43 species of Capsicum (including the domesticates), with new species still awaiting discovery in herbaria and in the field (especially in Peru and Bolivia).

Members of Capsicum plants are erect (e.g. C. schottianum, C. geminifolium), compact (e.g. C. chacoense, C. annuum var. annuum) or somewhat prostrate (e.g. C. annuum var. glabriusculum). They are subshrubs or shrubs, rarely trees (e.g. C. rhomboideum), short-lived perennials (e.g. C. chinense) or annual herbs (e.g. C. annuum var. annuum). Capsicum coccineum is unusual in being sprawling vines or scrambling shrubs. Stems are woody at the base (1.5–2.5 cm in diameter, rarely more) and some species have fissured bark and lenticels (e.g. C. rhomboideum, C. hookerianum); young stems are angular, herbaceous, usually hollow and weak and, occasionally, somewhat scrambling, range from glabrous to densely pubescent and may have anthocyanin along their length. The nodes are inflated and commonly green or purple.

Capsicum plants have typical solanaceous sympodial growth, giving the stems a “zig-zag” appearance. Initially, the vegetative growth is monopodial and the first stem to emerge has 8–39 leaves (C. annuum cultivars, Dorland and Went 1947; Patel and Shah 1977; Bosland and Votava 2000; C. chacoense, Barboza, pers. obs.) before the onset of sympodial ramification and flowering (Fig. 2A); the alternate leaves are arranged in a 2/5 phyllotaxic spiral. The main stem ends in a solitary flower or in a flower and one or two leaves (Fig. 2B, C). The number of leaves in a sympodial unit can be unifoliate or difoliate. In difoliate sympodial units, the leaves are geminate with the leaf pair slightly dissimilar in shape or size (e.g. C. cardenasii) or markedly dissimilar (e.g. C. dimorphum, C. lycianthoides) in size and/or shape.

Figure 2. 

Plant development in C. chacoense A monopodial vegetative growth B first dichotomy of the main stem and start of sympodial growth C initial branching with three branches. Photos by G.E. Barboza.

Species of Capsicum have simple leaves that are generally membranous or less frequently coriaceous (e.g. C. hunzikerianum, C. longifolium, C. pereirae), concolourous to discolourous, ovate to elliptical, rarely lanceolate or narrowly elliptical (C. longifolium, C. carassense) in outline; in taxa with geminate leaves, the minor leaves can be orbicular and sessile (C. dimorphum, C. lycianthoides). Leaf margins are always entire, rarely slightly revolute (C. caballeroi, C. ceratocalyx, C. hunzikerianum) and the leaf base is asymmetric, attenuate or truncate and sometimes decurrent on to the petiole (C. piuranum, C. rabenii). Leaf apices are obtuse or acute to acuminate or long-acuminate in few species (e.g. C. benoistii, C. piuranum, C. hunzikerianum). Petioles are longer in the domesticated species and in the major leaves of geminate leaf pairs in wild species.

Leaf and petiole anatomy of Capsicum members were investigated in domesticated species (Idu and Ogbe 1997; Weryszko-Chmielewsk and Michałojć 2011; Dias et al. 2013; Wahua et al. 2014) and, more extensively, in 13 wild species from different biogeographic environments (Palchetti et al. 2014). Most of the studied species have amphistomatic leaves, but leaves are hypostomatic in some C. frutescens and C. annuum cultivars, C. caatingae, C. cornutum and C. geminifolium (Idu and Ogbe 1997; Palchetti et al. 2014). Anisocytic, anomocytic and hemiparacytic stomatal types were reported; this variation was observed within a same leaf, a common feature in Solanaceae (Metcalfe and Chalk 1957; Bessis and Guyot 1979; Karatela and Gill 1986). Palchetti et al. (2014) described leaf blades with dorsiventral mesophyll, sparse to abundant calcium oxalate crystals (druses, solitary crystals and/or crystal sands), bicollateral vascular bundles surrounded by parenchyma, fibres or collenchyma and petioles with arch- or U-shaped vascular bundles. Leaf epidermal variables and the internal structural differences of leaf and petiole were not significantly different in the 13 wild species from different environments (Palchetti et al. 2014), but some anatomical characters were useful for the identification (stomata position and types, type of trichomes, type of cells surrounding the vascular bundle in the leaf, presence/absence and abundance and types of crystals and others).

Trichomes in Capsicum are mostly eglandular and simple, although branched trichomes can also occur (e.g. C. longidentatum, C. rhomboideum and C. parvifolium). Simple trichomes are uniseriate and usually 1–11-celled (Fig. 3A–D, F, K, O), with slightly (Fig. 3F–H) to strongly (Fig. 3A–D, O) minutely warty cuticle (Dias et al. 2013; Palchetti et al. 2014). Trichomes can be straight (Fig. 3A, B, O) or curved (Fig. 3C, D), cylindrical (Fig. 3B, O) or conical (Fig. 3A, M), flexible or somewhat rigid, antrorse or spreading. The distal cell of the trichomes can be acute (Fig. 3A, D, F), obtuse (Fig. 3H) or strongly curved resulting in a hook-shaped trichome (Fig. 3K). Branched trichomes can be furcate (forked) (Fig. 3G, P) to many times branched (Fig. 3J, L, N, see also Barboza et al. 2011). Papillae (Fig. 3E, I) are common at the margin and the apex of the corolla lobes in most species.

Figure 3. 

Eglandular trichomes of Capsicum species A–D, F, H, K, M, O simple trichomes E, I papillae G, J, L, N, P branched trichomes. Scale bar: 50 μm (A, B, E, H–J); 100 μm (C, D, F, G, K–P).

Glandular trichomes are common in Capsicum species. In most species, glandular trichomes are simple, with short uni- or bicellular stalks and globose to ellipsoid multicellular heads (Fig. 4D, E, G, I). These are usually distributed on stems, leaves, pedicels and the outer calyx surface of some species (e.g. C. caatingae, C. cardenasii, C. galapagoense, C. tovarii) and on the inner calyx surface in all Capsicum species. Glandular trichomes with (1–) 2–3-celled stalks and globose or globose-peltate unicellular heads (Fig. 4B, C, F) are found on the interior surface of the corolla in many species. The most unusual glandular trichomes are found in C. eshbaughii (Fig. 4A, H, J) which has a distinctive branched glandular indument (Barboza 2011).

Figure 4. 

Glandular trichomes of Capsicum species. Scale bar: 50 μm (A–G, I); 100 μm (H, J).

Density of pubescence is highly variable both within and between species. Sometimes, this variability has been considered diagnostic at the infraspecific level. For example, the densely pubescent plants of C. chacoense, C. baccatum, C. eshbaughii or C. annuum have been recognised at the infraspecific level (see descriptions and synonymy of those species).

Capsicum species have axillary flowers; they are solitary only in C. chacoense (Fig. 2C) or may be in 1–2 (–3)-flowered inflorescences in some species (e.g. C. baccatum, C. friburgense, C. piuranum) or in 4–13-flowered inflorescences in most of the species. Capsicum ceratocalyx and C. caatingae may have more than 18 flowers per inflorescence. The inflorescences are either epedunculate and unbranched or borne on a short (e.g. C. ceratocalyx, C. coccineum) or somewhat elongate rachis that is occasionally forked (e.g. C. regale). Flowers are usually deciduous, falling off the plant gradually and leaving conspicuous or obscure pedicel scars on the rachis.

All Capsicum flowers have distinct, usually pubescent, pedicels that may be terete (Fig. 5B), angled (Fig. 5A) or, rarely, almost winged (C. ceratocalyx). Pedicel/flower position is a useful identification character. Pedicels (and flowers) can be deflexed (termed pendent in this treatment; for example, C. pereirae, C. lanceolatum, Fig. 5C) or erect to spreading. In some cases, the pedicels are distally abruptly twisted forming a right or acute angle with the longitudinal axis of the flower (termed geniculate in this treatment). Flowers with geniculate pedicels may be orientated with their longitudinal axis in horizontal position (e.g. C. rabenii, Fig. 5A) or nearly parallel to the pedicel axis (e.g. C. annuum var. glabriusculum, Fig. 5B). In fruits, pedicels are usually green (Fig. 6A, B, G) or green with purple stripes (e.g. C. eximium, C. longifolium) or completely purple (C. regale, Fig. 6F). They are normally widened distally and can be pendent (e.g. C. recurvatum, C. flexuosum) or erect (e.g., C. baccatum var. baccatum, C. chacoense, C. frutescens).

Figure 5. 

Flower morphology in Capsicum species A C. rabenii B C. annuum var. glabriusculum C C. lanceolatum D C. schottianum E C. frutescens F C. eximium G C. eshbaughii H C. cornutum I C. galapagoense J C. recurvatum K C. cardenasii L C. lycianthoides M C. chacoense N C. baccatum var. baccatum O C. caballeroi. Abbreviations. im interpetalar membrane sp staminal plaque.

Figure 6. 

A–G Fruit morphology in Capsicum species H, J epicarp structure A C. chinense B C. baccatum var. baccatum C C. hookerianum D C. lanceolatum E C. coccineum F C. regale G C. schottianum H epicarp with regular epidermal cells I epicarp with some sclereids amongst the regular epidermal cells J epicarp exclusively with sclereids. Abbreviation. sc sclereids. Scale bar: 10 μm (H, I, J).

The calyx in Capsicum species is usually 5-merous (4–8-merous in domesticated taxa) and entirely synsepalous (Fig. 5E). The calyx tube is generally cup-shaped or campanulate and 5–10 nerved, with the margin always entire (Fig. 5E, L). Below the margin, up to10 calyx appendages may emerge (Fig. 5A, F–H, L), a character only shared with Lycianthes (Dunal 1852; Bitter 1919; D’Arcy 1986; Hunziker 2001). In calyces without appendages, the calyx outline can be circular with the five main veins completely immersed in the calyx tube and ending very near its margin (e.g. C. campylopodium, C. galapagoense) (Fig. 5E). In species such as C. schottianum, the calyx outline may be pentagonal (Fig. 5D) with the main veins ending in five tiny lateral umbos or lumps that barely emerge over the calyx margin. Calyx appendages are lateral elongate expansions of the calyx tube (Fig. 5C, F–H, L) with vasculature formed by each main or secondary vein arching outwards into the appendages and returning to the calyx tube to end in the margin of the calyx (see figs 1–4 in D’Arcy 1986). The calyx sleeve, which is a ring of tissue immediately below the margin, is rudimentary (e.g. C. recurvatum, C. hookerianum, C. coccineum, C. lycianthoides, Fig. 5L) or non-existent in Capsicum (Fig. 5F, H). This sleeve is well developed in many Lycianthes species (D’Arcy 1986; Dean et al. 2017, 2020) and, thus, serves as a trait to differentiate the two genera. The number, shape, size and orientation of the appendages are important identification characters within Capsicum. The number of appendages is consistent (5 or 10) in most species, but in others, it can vary (2–10) within a species and, sometimes, even within an individual (e.g. C. recurvatum). Appendages shape ranges from subulate (wider at the base and pointed at the tip, for example, C. lycianthoides, Fig. 5L), to cylindrical (e.g. C. eximium, Fig. 5C, F) and linear (e.g. C. eshbaughii, C. cornutum, Fig. 5G, H). In some species, the appendages are thick and strongly laterally flattened like triangular-compressed wings (e.g. C. longifolium, C. regale). Appendages are usually equal or subequal (e.g. C. eximium, C. piuranum), but when there are more than five in number, the main appendages (those with vasculature formed by the main veins) are longer than the secondary appendages (those with vasculature formed by secondary veins) (e.g. C. hookerianum, C. longidentatum). The longest main appendages (ca. 8.5 mm) are found in C. cornutum (Fig. 5H). Calyx appendages can be erect and appressed to the corolla (e.g. C. eximium, Fig. 5F), spreading (e.g. C. dimorphum, C. lycianthoides, Fig. 5L), recurved (e.g. C. recurvatum), reflexed (e.g. C. lanceolatum, Fig. 5C) or incurved and horn-like (C. ceratocalyx); orientation may change in the transition between flowering and fruiting (e.g. C. coccineum). An annular constriction at the junction between the fruiting calyx and pedicel is present in a few wild species (e.g. C. caatingae, C. regale) and it is a key character used in the identification of C. chinense (Fig. 6A). Fruiting calyces are persistent and not accrescent or only slightly accrescent (domesticated species). They are discoid in most species (Fig. 6G), but can be cup-shaped (e.g. C. frutescens) and are strongly reflexed in C. coccineum (Fig. 6E).

Capsicum species have 5-merous (6–8-merous in domesticated taxa) sympetalous corollas. Corollas are usually of intermediate size (6–14 mm long), the smallest measuring 4–5 mm long (e.g. C. galapagoense) and the largest ones reaching 17–18 mm long (e.g. C. caballeroi, C. piuranum). Most species have stellate corollas (Fig. 5D–G, I, J), but campanulate (e.g. C. lanceolatum, Fig. 5C, C. cardenasii, Fig. 5K), broadly campanulate (C. lycianthoides, Fig. 5L, C. rhomboideum), campanulate-urceolate (C. friburgense), rotate-stellate (C. flexuosum, C. baccatum var. baccatum, Fig. 5N) or rotate (e.g. C. pubescens) corollas are also present. The corolla tube can be very short (C. benoistii), short (e.g. C. eximium, Fig. 5F) or long (e.g. C. caballeroi, C. cardenasii, Fig. 5K, C. piuranum). The lobes are connected by a thinner interpetalar membrane nearly completely (e.g. C. lycianthoides, Fig. 5K, L), partly (e.g. C. baccatum var. baccatum Fig. 5J) or just near the base (C. annuum var. glabriusculum, C. galapagoense, Fig. 5I); in very few species, interpetalar membrane is absent (e.g. C. caballeroi).

Corolla colour is highly variable in Capsicum species. Corollas can be entirely white (e.g. C. chacoense, C. galapagoense, C. annuum var. annuum), dull white or greenish-white (C. chinense, C. frutescens), light yellow (C. neei), yellow (e.g. C. piuranum, C. caballeroi) or violet or fuchsia (C. friburgense). Other species have corollas with a predominant primary colour (white, yellow or purple), as well as markings (spots) with diverse pigmentation. The abaxial surface (outer surface) of the corollas may have the same colouration as the adaxial surface (inner surface) (e.g. C. geminifolium, C. regale) or it may have a faded or a different colouration (e.g. C. tovarii, C. pereirae). In many species, the co-occurrence of different pigments results in multi-coloured corollas, for example, white corollas with purple (or variations) spots at the base of the lobes and limb and green or greenish-yellow centre (e.g. C. villosum, C. schottianum, C. pereirae). Descriptions in literature or on specimen labels usually refer to the colour of the inner (adaxial) corolla surface. In this monograph, description of the corolla colour of both surfaces is provided for each species; corolla colour can be very difficult to see on herbarium specimens.

Corolla pigmentation is due to anthocyanins which produce violet or purple shades (e.g. C. lycianthoides, C. lanceolatum, Fig. 5C, F, K, L) and chlorophyll which produces green colouring (C. baccatum, C. flexuosum, Fig. 5D, J).

The adaxial surfaces of the corollas are glabrous (many Andean species) or may be covered by sparse glandular trichomes (e.g. C. ceratocalyx, C. tovarii) or have a continuous ring of glandular trichomes (Fig. 4C, F) at the base of the lobes and in the throat (the Bolivian and Brazilian species). The abaxial surfaces of corolla lobes are mostly papillate, with minutely warty papillae (Fig. 3E, I) or with short simple trichomes on the margins and tips. The lobes are spreading in most species (Fig. 5D, F, G, I, J), erect in C. tovarii or slightly (e.g. C. cardenasii, Fig. 5K) to strongly recurved in some species (C. caballeroi and C. friburgense).

Capsicum species have usually five (sometimes 6–8 in domesticated taxa) equal stamens; unequal stamens have only been observed in three species: C. campylopodium (and sometimes also C. lycianthoides), which has two stamens longer than the other three (Hunziker 2001) and C. regale, which has one longer than the other four (Barboza et al. 2020b). The free portion of the filaments is always distinct and glabrous and it is usually longer than the anthers (Fig. 5N). Each filament broadens at its base forming a staminal plaque (called a stapet, Hunziker 2001), with two short lateral auricles fused to the corolla base. In C. chacoense, these lateral auricles are long and are not fused to the corolla. The anthers are longitudinally dehiscent and are usually ellipsoid and yellow or cream in colour (Fig. 5M); in domesticated taxa and some wild species (e.g. C. dimorphum, C. carassense, C. regale), the anthers are blue (Fig. 5B), bluish-grey or purple. In pre-anthesis and early anthesis, the anthers are connivent (Fig. 5M), but they become separated during anthesis (Fig. 5N), remaining somewhat connivent in some species (Fig. 5B). Anther size is variable, ranging from 0.9–1.3 mm long in C. galapagoense, to 1–3 mm long in most species, to up to 4 mm in C. hunzikerianum.

Pollen is yellow or white, trizonocolporate, spheroidal, prolate, prolate-spheroidal or oblate-spheroidal, with a triangular to circular outline in polar view. It is usually small, from 15 µm in C. rhomboideum (Bo and Carrizo García 2015) to 18–24.73 µm in some domesticated species (Dharamadhaj and Prakash 1978; Martins et al. 2013; Adedeji and Akinniyi 2015; Bo and Carrizo García 2015; Kayani et al. 2018; Yang et al. 2018; Song et al. 2019), but in other domesticated and wild species, sizes range from 25.75 to 38 µm (Murray and Eshbaugh 1971; Quagliotti 1979; Martins et al. 2013). Pollen grains are shed in monads and at the binucleate stage (Cochran 1938; Yaqub and Smith 1971; Dharamadhaj and Prakash 1978; Kim et al. 2004; Bo and Carrizo García 2015) or trinucleate stage (e.g. a C. frutescens cultivar, Lengel 1960). Information on the ornamentation of the exine is variable in literature depending on whether observations were made with a light microscope (LM) or a scanning electron microscope (SEM). The exine appears to be clearly microechinate and perforate under SEM in C. annuum (Halbritter 2016a; Song et al. 2019) and C. pubescens (Bo and Carrizo García 2015; Halbritter 2016b). The exine has been described as reticulate (Murry and Eshbaugh 1971; Roubik and Moreno 1991), foveolate (Martins et al. 2013), psilate or faintly granulate (Kayani et al. 2018) and scabrate (Barth and Duarte 2008), based on LM observations for some Capsicum species, which should be corroborated with SEM. Pollen morphology is lacking for most wild Capsicum species and more information is needed to evaluate the usefulness of pollen to distinguish species.

The gynoecium in Capsicum species is usually bicarpellate (2–5-carpellate in domesticated species). The ovary is superior with axile placentation, glabrous and usually subglobose to ovoid, rarely ellipsoid (e.g. C. frutescens). The style is simple, straight or slightly curved, cylindrical (the same width from the proximal to distal end, Fig. 5N) or clavate (broadened gradually from its base to the apex), glabrous, white, cream, lilac or purple and commonly exserted beyond the anthers. Most species have homostylous flowers, but some species (C. annuum, C. baccatum, C. benoistii, C. tovarii, C. pubescens) have heterostylous flowers, with different flowers on the same plant bearing short, medium or long styles. In domesticated species, style length variations have been observed amongst cultivars (Bosland and Votava 2000; Peña-Yam et al. 2019). The stigma is pale green or cream, globose or discoid, sometimes slightly bilobed (e.g. C. longifolium, C. lanceolatum) and finely papillate. The ovules are usually numerous, anatropous, tenuinucellate, with a single integument (Cochran 1938; Munting 1974; Dharamadhaj and Prakash 1978; Pérez-Pastrana et al. 2018). The nectary is located at the base of the ovary and is easily observed in fresh material, but sometimes can be very difficult to see in herbarium specimens; it is an inconspicuous annular disc, paler in colour than the rest of the ovary, variable in thickness and produces copious nectar. The nectar is colourless and is exposed in five nectar droplets on the corolla limb (see more detail in Floral biology and pollination).

The fruit is usually a bicarpellate berry (Fig. 7A) that can also be 3–5-carpellate in domesticated species (e.g. C. annuum var. annuum, C. chinense, C. pubescens) or some wild species (e.g. C. tovarii). The berries are extremely diverse in shape, size and colour in domesticated species and more homogeneous in wild (and semi-domesticated) species. The fruits are juicy, fleshy, opaque (most species) or translucent (some Brazilian species). Wild species have globose or subglobose berries < 15 mm in diameter (Fig. 6A, C, E–G) or short ellipsoid or ovoid (e.g. C. baccatum var. baccatum, C. chacoense) berries 0.6–14 mm long (Fig. 6B). The fruits in domesticated species are much longer and wider, with the most elongate fruits (300 mm long or more) in C. annuum var. annuum (Bosland and Votava 2000). The great variation of the fruit shapes has resulted in the proposal of more than 100 names applied to the domesticated species (here assigned to any of the five recognised domesticated taxa). Within each of the domesticated taxon, a system of different fruit types (i.e. characterised by a defined fruit shape and colour, pungency level, aroma and/or flavour and uses) is used by horticulturists or plant breeders (Bosland et al. 1988); nearly 100 fruit types are known, the most frequent of which are described in Bosland and Votava (2000) and DeWitt and Bosland (1997, 2009). Red is the most frequent fruit colour in wild Capsicum species (Fig. 6B–E), with colours ranging from orange-red (e.g. C. geminifolium), dark burgundy (C. rhomboideum) to dark blue or purple-blue (C. regale, Fig. 6F). Some Brazilian species have greenish-golden yellow, translucent fruits (e.g. C. parvifolium, C. schottianum, Fig. 6G), a character state considered to be derived within Capsicum (Carrizo García et al. 2016).

Figure 7. 

Fruit anatomy in Capsicum species A, D–F C. baccatum var. pendulum B C. baccatum var. umbilicatum C, G, H C. pubescens A fruit, in cross section (note giant cells in the pericarp) B one locule of a fruit, in cross section (note the absence of giant cells in the pericarp) C, D epicarp and some layers of mesocarp (in D, observe cuticular wedges) E sector of pericarp (the arrow indicates the increase of the cell size ending in the giant cells) F detail of two adjacent giant cells G sector of homogeneous endocarp H sclereids of the endocarp. Abbreviations. c cuticle, cw cuticular wedge, epc epidermal cells, gc giant cells, p pericarp. Scale bars: 1 mm (A, B, E); 10 μm (C, D, H); 100 μm (F, G).

Fruiting pedicels are usually green (Fig. 6A, B, G) or green with purple stripes (e.g. C. eximium, C. longifolium) or completely purple (C. regale, Fig. 6F). They are normally widened distally and can be pendent (e.g. C. recurvatum, C. flexuosum) or erect (e.g., C. baccatum var. baccatum, C. chacoense, C. frutescens). Pedicels and ripe fruits are persistent in domesticated species, staying attached to the plant; in most wild species, the mature berries are usually deciduous and are easily detached from the calyx, leaving only pedicels and fruiting calyces on the plant (Carvalho et al. 2014, GEB, pers. obs.). In just a few species (e.g. C. muticum, C. coccineum), mature berries fall from the plant with pedicels attached, leaving conspicuous scars.

The development of the pericarp in Capsicum species is the typical of a true berry (Roth 1977), with the epicarp and endocarp originating respectively from the outer and inner epidermis of the ovary wall and the mesocarp derived from the middle layers between the two epidermal layers (Roth 1977; Filippa and Bernardello 1992); this structure is typical of most Solanaceae (Roth 1977; Bernardello 1983; Filippa and Bernardello 1992). The taxonomic significance of the pericarp structure in Capsicum has been rarely considered with some observations made on varietals or cultivars of the domesticated species (Tschirch and Oesterle 1900; Augustin 1907; Tschirch 1925; Winton and Winton 1939; András 1966; Fridvalszky and Nagy 1966; Konecsni 1971; Munting 1974; Dave et al. 1979; Dave 1986; Weryszko-Chmielewska and Michałojć 2011; Dias et al. 2013) and a few observations made on wild species (Filippa and Bernardello 1992). In this monograph, we include the most relevant taxonomic characters observed in nearly all the species of Capsicum (see Suppl. material 1: Appendix 1, for full details of each species).

The epicarp consists of a uniseriate epidermis covered by a smooth (e.g. C. annuum, C. chacoense, C. pubescens, Fig. 7C) or striate cuticle (e.g. C. campylopodium, C. mirabile). The cuticle can be thin (4.5–11 µm, for example, C. flexuosum, C. campylopodium) or thick (11.5–19 µm, for example, C. eximium, C. galapagoense) and usually projects towards the anticlinal walls of two adjacent epidermal cells forming conspicuous cuticular wedges (Fig. 7D). The cuticular wedges can be shallow to very deep, reaching the first layer of the mesocarp, with the cuticle extending below the inner periclinal wall of the epidermal cells (e.g. C. caatingae); these cuticular wedges vary from 7.5 to 54 µm thick. The epicarp is formed by regular epidermal cells (Fig. 6H, I), scarce stomata and sometimes sclereids (Fig. 6I, J). In cross section, epidermal cells are elongate tangentially (rectangular, Fig. 7C, D), isodiametric or flask-shaped (e.g. C. hookerianum) and in superficial view, they are polygonal, with straight (Fig. 6H) or slightly sinuous cell walls; pits appear frequently in thick- and thin-walled cells (Fig. 6I). The epicarp may have scattered sclereids amongst the epidermal cells (e.g. C. baccatum var. pendulum, Fig. 6I) or it can consist exclusively of sclereids (C. baccatum var. baccatum, Fig. 6J). Ventilating clefts are found in the epicarp of many species (Dave et al. 1979; Filippa and Bernardello 1982; this work), but their presence is not a constant feature within a species (Dave 1986).

The mesocarp consists of (5–) 7–26 layers, with up to 30 layers found in the thick pericarp of a sweet C. annuum var. annuum cultivar known as “calahorra”. The mesocarp can be homogeneous or heterogeneous; a homogeneous mesocarp is formed exclusively by parenchyma (thin-walled cells) or collenchyma (thick-walled cells) (e.g. C. rhomboideum, C. hookerianum), whilst a heterogeneous mesocarp consists of both collenchyma and parenchyma with a variable number of layers for each tissue, depending on the species (see Suppl. material 1: Appendix 1). The collenchymatous layers are always placed underneath the epicarp and both together have been referred as the “exocarp” (Roth 1977). The parenchymatous layers are underlying the collenchyma and the number of layers is usually greater than those of the collenchyma. Crystal sand and vascular bundles are found in the collenchyma or parenchyma (e.g. C. rhomboideum, C. dimorphum). A gradual increase of cell size occurs from the periphery inwards (Fig. 7E). In a few Capsicum species (e.g. the Andean clade species), the cells of the innermost layer of the mesocarp are equal in size and similar in shape to the adjacent mesocarp cells (Fig. 7B); thus, the pericarp surface facing the locule (endocarp) is smooth. In contrast, in most species, the inner surface of the pericarp is blistered due to the extensive development of the inner hypodermis (Fig. 7A, E, F); the inner hypodermis develops as a single layer of ‘giant cells’ that are initially large in the ovary wall (Augustin 1907; Munting 1974) and then increase tremendously in size towards fruit maturity (Fig. 7A, E). These giant cells are occupied by scanty cytoplasm, a large watery vacuole and a large nucleus pushed to the periphery by the vacuole. The size of the giant cells varies amongst the species; they can be nearly as long as wide, ranging from 390–1380 µm long and 300–1050 µm wide (most species) or 2–6 times longer than wide, varying from 540–2500 µm long and 150–490 µm wide (e.g. some Brazilian species). Each giant cell is attached to its neighbouring cell and the endocarp and they are surrounded by a triangular multiseriate wedge-shaped cluster of parenchymatous cells between their anticlinal cell walls (“bridge”, fide Roth 1977). The development of giant cells in the mesocarp is a derived anatomical trait in Capsicum, absent in the Andean clade and with a single reversion to the ancestral state in C. baccatum var. umbilicatum (Fig. 7B) (Carrizo García et al. 2016).

Hard inclusions of sclereids (stone cells) are developed in the mesocarp of some species (5 spp., see Suppl. material 1: Appendix 1); the stone cells can be completely immersed in the mesocarp and then only visible under microscope (C. piuranum) or they are easily seen in the pericarp surface in dried fruits (e.g. C. geminifolium). There are never more than six per fruit. The presence, number and position of the stone cells are not consistent within a single species; thus, these are not useful traits in the identification of Capsicum species, as they may be in other Solanaceous genera, such as Lycianthes (Bitter 1919) or Solanum (Särkinen et al. 2018; Knapp et al. 2019).

The endocarp develops from the inner epidermis of the ovary wall; it consists of one layer of polygonal or irregular cells (surface view) with straight or sinuate cell walls. The endocarp can be homogeneous, that is entirely with thin (e.g. C. recurvatum) or pitted thick-walled cells (sclereids) (e.g. C. rhomboideum, C. pubescens, Fig. 7G, H) or it may consist of a mixture of cells with thin and thick walls forming a heterogeneous tissue (e.g. C. annuum); in this latter endocarp type, groups of pitted sclereids lie on the inner periclinal walls of the giant cells that alternate with groups of thin-walled cells placed on the multiseriate wedge (“bridge”) of parenchymatous cells (Konecsni 1971; Dave 1986).

Some structural features of the pericarp useful in species-level taxonomy are detailed in Suppl. material 1: Appendix 1.

The presence of the pungent principles (capsaicinoids) of Capsicum fruits within cells of the interlocular septum and placenta has been demonstrated by histochemical (Ohta 1962d) and radioisotopic (Iwai et al. 1979) evidence, structural and ultrastructural microscopic analyses (Suzuki et al. 1980; Zamski et al. 1987; Stewart et al. 2007) and analytical chemical procedures (Manirakiza et al. 2003; Stewart et al. 2007; Nugroho 2016; Guillen et al. 2018). The histological characteristics of the septum during fruit development have been investigated extensively in some cultivars of C. annuum and C. frutescens (Tschirch and Oesterle 1900; Ohta 1962d; Munting 1974; Dave et al. 1979; Suzuki et al. 1980), as well as some wild Capsicum species (Filippa and Bernardello 1992). In general, the septum consists of epidermal cells and a loose parenchymatous tissue with intercellular spaces. The epidermal cells are responsible for the synthesis, accumulation (in pockets or blisters) and secretion of capsaicinoids. In early stages of fruit development (about 10 days after flowering), some of these cells become secretory (Ohta 1962d; Suzuki et al. 1980; Filippa and Bernardello 1992), but they fully develop the characteristic traits of a glandular tissue (i.e. large elongate size, abundant cytoplasm, a large vacuole, many small vesicles containing electron-dense granules and a large nucleus) nearly 30 days after flowering (Suzuki et al. 1980).

An ultrastructural study demonstrated that the major reservoir of capsaicinoids is in the capsisome, a specific capsaicinoid biosynthesising and accumulating vacuole, different from the vacuoles regarded as reservoirs of organic acids (Fujiwake et al. 1980, 1982).

Capsaicinoids have also been found in the pericarp and seeds in significant or low amounts in some species and cultivars (e.g. C. chinense, C. baccatum) (Pandhair and Sharma 2008; Bosland et al. 2015; Guillen et al. 2018).

The gross morphology of the seeds and details of the sculpturing of the seed coat for all Capsicum species are summarised in Suppl. material 2: Appendix 2. The attributes used in species descriptions are illustrated in Fig. 8.

Figure 8. 

Seed morphology and seed coat structure A Capsicum chinense B C. chacoense, longitudinal section C, D C. schottianum, cross section (D detail of the seed coat structure) E C. annuum var. annuum, detail of seed coat structure (the rectangle indicates a cell of the seed coat) F, G C. eximium, cross sections at the seed margin and seed body, respectively H C. dimorphum, cross section at the seed body. Abbreviations. aw anticlinal cell wall, bp beak prominence, em embryo, en endosperm, h hilum, im inferior seed margin, ipw inner periclinal wall, opw outer periclinal wall, sb seed body, sc seed coat, sm superior seed margin, w seed width, l seed length. Scale bars: 200 μm (A–C); 100 μm (D, F, H); 20 μm (E, G).

Seed shape (and size) is influenced by the position in the berry. The seeds are flattened to slightly angled, mostly C- or D-shaped (Gunn and Gaffney 1974), subglobose or ellipsoid, rarely reniform or teardrop-shaped. Seed shape is sometimes difficult to define in the domesticated species due to the presence of a protrusion on the seed coat (a beak-like prominence) in a vertical or nearly vertical direction above the hilum (Fig. 8A, B). This protrusion can also occur in a lateral position in some wild species with the hilum placed on the protrusion edge (e.g. C. cornutum). Seed size varies from small (1.5–2.5 mm long) and medium (2.6–3.9 mm long) to large (4–7 mm long). Capsicum lycianthoides has the smallest seeds (1.5–1.8 mm long), whereas the largest seeds are found in the domesticated C. pubescens (5.5–7 mm long). Seed colour is usually uniform and is slightly shiny to shiny when observed dry under a stereomicroscope. Seeds can be grouped in three categories based on their colour: (1) pale yellow or nearly white (C. neei) to yellow (mostly domesticated species and their close relatives); (2) brownish-yellow to brown (many Bolivian species); and (3) brownish-black to black (many Andean and Brazilian species). Seed number per berry in Capsicum species is usually 10–45, with only four seeds found in C. campylopodium and more than 50 in C. lanceolatum, C. piuranum and some domesticated taxa.

The hilum is always marginal (on the inferior margin, Fig. 8A) and its position may be medial (Fig. 9Q), subterminal (Figs 9N, 10N) or terminal (Fig. 9I). The hilum area may be inconspicuous (minute to small) or conspicuous (medium to large). The hilum area shape may be elliptical, linear, ovoid or more rarely triangular (C. minutiflorum, C. pubescens) or circular (C. tovarii); some species may have two or three different hilar area shapes.

Figure 9. 

Seeds and seed coat morphology in species of the Annuum Clade A–D C. annuum var. annuum E–H C. annuum var. glabriusculum I–L C. frutescens M–P C. chinense Q–T C. galapagoense A, I seeds with testa partly digested B seed coat with the external periclinal cell wall partly removed C, D cross section of the seed at the seed margin and seed body, respectively; E, M, N, Q untreated seeds showing subterminal hilum (E, N) and medial hilum (Q) F, J, O, P, S, T detail of a non-digested portion of the seed coat G, K testa pattern with the external periclinal cell wall removed H, L detail of testa cells R hilum. Abbreviation. opw outer periclinal cell wall. Scale bars: 200 μm (A, E, I, M, N, Q); 20 μm (B, F–H, K, L, P, S, T); 50 μm (C, J, O, R); 10 μm (D).

Figure 10. 

Seeds and seed coat morphology in species of the Baccatum Clade A–D C. baccatum var. baccatum E C. baccatum var. pendulum F–H C. baccatum var. umbilicatum I–L C. chacoense M–O C. rabenii. A, I, M seeds with testa partly digested B, G detail of a non-digested portion of the seed coat C, H, J, K, O testa pattern of treated seeds showing anticlinal cell walls with fibrils (C, K), papillae (H) and ridge (O) D detail of a testa cell E, F untreated seeds L cross section of the seed at the seed body N seed showing the subterminal elliptical hilum. Scale bars: 10 μm (H); 20 μm (B–D, G, K, L, O); 100 μm (J); 200 μm (A, E, F, I, M, N).

Capsicum annuum (and its varieties) has been the most frequently studied species (Lohde 1875; Moeller 1886, 1905; Hanausek 1888; Hartwich 1894; Wojciechowska 1972; Krstić et al. 2001, Kong et al. 2011) before our studies reported here. Some other authors focused on the development and the structure and sculpture of the seed coat of many Solanaceae species, including some of the domesticated capsicums (Souèges 1907; Gunn and Gaffney 1974; Al-Nowaihi and Mourad 1999; Dias et al. 2013). More recently, Chiou and Hastorf (2014) provided a morphometric approach, based on 27 qualitative and quantitative seed attributes for the five domesticated species. They recovered some useful traits that can be used to compare features of modern Capsicum seeds to archaeological seeds. The gross morphology and the sculpture of the seed coat provided here for each Capsicum species (Suppl. material 2: Appendix 2) highlight the importance of some characters for delimiting species, species groups or clades. Carrizo García et al. (2016) found that seed colour is useful for identifying species or small clades (e.g. brownish-black or black seeds in the Atlantic Forest and in Andean clades or the pale yellow seeds in the Annuum and Baccatum clades). The seeds of species within the same clade are also characterised by similar sculpture and the structure of the outer layer. For example, species of the Atlantic Forest clade all have a seed coat that is reticulate-tuberculate/reticulate with pillar-like structures, has deep cells in the margin and seed body and has a thin cellulosic outer periclinal cell wall in the outer layer that is easily removed (naturally or by enzyme etching).

Seed ornamentation refers to the appearance of the seed coat (testa) (Figs 9–18). The testa appearance varies in the majority of the species, depending on the magnification (naked eye, stereomicroscope or SEM) and whether or not the seeds are untreated or treated with a technique, such as enzyme etching. By removing the outer periclinal wall with enzyme etching, the ornamentation of the seed coat (sculpture) can be observed in detail under SEM. From the SEM data obtained of all treated seeds, the sculpture of the seed coat and variations in the anticlinal (lateral) walls of the outer epidermal layer vary between species and provide useful diagnostic features.

Five major types of seed coat sculpture were observed after the outer periclinal wall was removed by enzymatic digestion: (1) reticulate, with straight to wavy cell walls (Fig. 14B, C, M, O, for example, C. dimorphum); (2) cerebelloid, with sinuate to strongly sinuate cell walls (Fig. 15B, E, F, K, for example, C. hookerianum); (3) reticulate-cerebelloid, reticulate at the seed margins and cerebelloid in the seed body (Fig. 16A, B, for example, C. caballeroi); (4) reticulate with pillar-like outgrowths at margins (Fig. 11M, R, S, for example, C. cornutum, C. mirabile), these outgrowths due to unequal height of the anticlinal walls thickenings (Fig. 8C, D); and (5) cerebelloid with pillar-like outgrowths at margins (Fig. 11B, C, for example, C. regale and some seeds of C. campylopodium and C. schottianum). The most common combinations in the ornamentation of untreated/treated seeds in each species are: smooth to obscurely reticulate/cerebelloid (Figs 9I, K, 10A, C); reticulate, marginally tuberculate/reticulate with pillar-like outgrowths at margins (Figs 11I–K, Q–S, 12L–O, 13E–H); reticulate/reticulate (Fig. 14A–C); smooth and reticulate at margins or completely reticulate/reticulate-cerebelloid (Fig. 15I–K) (see Suppl. material 2: Appendix 2 for each species).

Figure 11. 

Seeds and seed coat morphology in species of the Atlantic Forest Clade A–D C. campylopodium E–H C. carassense I–L C. cornutum M–P C. friburgense Q–T C. mirabile A untreated seed B, I, Q seeds with testa partly digested C, F, K, O, S marginal testa pattern of treated seeds (note pillar-like outgrowth in K, O, S) D, G, H, L, P, T testa pattern at the seed body of treated seeds showing anticlinal cell walls papillate and punctate E, J, M, R treated seeds (in J showing lateral prominence) N hilar zone with a linear hilum. Abbreviation. lp lateral prominence. Scale bars: 200 μm (A, B, E, I, J, M, Q, R); 100 μm (C, G, K, N, O, S); 20 μm (D, F, H, L, P, T).

Figure 12. 

Seeds and seed coat morphology in species of the Atlantic Forest Clade A–C C. muticum D–G C. pereirae H–K C. mirum L–P C. schottianum Q–T C. recurvatum A, E, I, N, Q treated seeds B, F, J, O, S marginal testa pattern of treated seeds with pillar-like outgrowths C, G, P, T testa pattern at the seed body of treated seeds showing anticlinal cell walls papillate and punctate D, H, L untreated seeds K detail of papillate anticlinal cell walls M detail of a non-digested portion of the seed coat R hilar zone with a linear hilum. Scale bars: 200 μm (A, D, E, H, I, L, N, Q); 100 μm (B, F, J, O, R); 20 μm (C, G, K, M, P, S, T).

Figure 13. 

Seeds and seed coat morphology in species of the Atlantic Forest Clade A–D C. hunzikerianum E–I C. villosum A, G seeds with testa digested B seed showing medial and linear hilum C, H marginal testa pattern of treated seeds D, I testa pattern at the seed body of treated seeds showing anticlinal cell walls papillate (D) and punctate (I) E untreated seed F detail of a non-digested portion of the seed coat. Scale bars: 200 μm (A, B, E, G); 20 μm (C, D, F, I); 100 μm (H).

Figure 14. 

Seeds and seed coat morphology in species of the Andean Clade A–D C. dimorphum E–H C. geminifolium I–L C. lanceolatum M–P C. longifolium Q–T C. lycianthoides A, Q, S seeds untreated B, E, I, J, M seeds with testa digested C, G, K, O testa pattern at the seed body of treated seeds showing anticlinal cell walls punctate (C, J, O) and with fibrils (G, K, O) D detail of ridge H, L, P detail of testa cells F, N hilar zone R, T detail of a non-digested portion of the seed coat. Scale bars: 200 μm (A, B, E, I, J, M, Q, S); 100 μm (C, K, N, O); 20 μm (D, F, G, H, L, P, R, T).

Figure 15. 

Seeds and seed coat morphology in species of the Andean Clade A–D C. hookerianum E–H C. piuranum I–K C. rhomboideum A untreated seed B, E treated seeds C, D, G, J, K testa pattern of treated seeds showing anticlinal cell walls punctate (C, D, J, K) and papillate (G) F hilar zone H detail of testa cells densely papillate I seed partly digested. Scale bars: 200 μm (A, B, E, I); 20 μm (C, D, F, G, H, J, K).

The anticlinal cell walls observed with SEM are either: (1) papillate, with papillae 2.5–4 µm in diameter on the cell walls (Figs 10H, 11D, H, T, 12C, G, K, T, 15H, 16D, H, L, P, T, 17H, L, O, 18D, H, L); and (2) punctate, with perforations (holes, ca. 35 µm in diameter) at the bottom (Figs 9L, 11G, H, L, T, 12C, G, 14L, 16S) or uniformly distributed on the cell walls (Figs 11O, P, 12P, T, 14P, 15D, K, 17G, N).

Figure 16. 

Seeds and seed coat morphology in species of the Bolivian Clade A–D C. caballeroi E–H C. ceratocalyx I–L C. minutiflorum M–P C. neei Q–T C. coccineum A, Q seeds with testa partly digested B, F, G, J, O, R marginal testa pattern of treated seeds showing anticlinal cell walls punctate and densely papillate (G, O), punctate (J) and with ridge (R) C, H, K, P, S detail of testa cells D, L, T detail of papillae E, I, M treated seeds N linear hilum. Scale bars: 300 μm (A, E, I, M, N, Q); 50 μm (B, F, G, H, O, P); 20 μm (C, J, K, L, R, S); 5 μm (D, T).

Figure 17. 

Seeds and seed coat morphology in species of the Caatinga, Longidentatum, Flexuosum and Tovarii Clades A–D C. caatingae E–H C. parvifolium I–L C. longidentatum M–O C. flexuosum P–R C. tovarii A, M, P seeds with testa partly digested (in A hilum in terminal position, P hilum subterminal) B detail of a non-digested portion of the seed coat C, F, J, N, Q marginal testa pattern of treated seeds showing anticlinal cell walls papillate (F), with fibrils (J), punctate (N) and with fringe (Q) D, G, K, O, R detail of testa cells E, I treated seeds H, L papillae on anticlinal cell walls Scale bars: 200 μm (A, E, I, M, P); 20 μm (B, D, G, H, J, K, L, N, O, Q, R); 100 μm (C, F).

Figure 18. 

Seeds and seed coat morphology in species of the Purple corolla and Pubescens Clades A–D C. cardenasii E–H C. eshbaughii I–L C. eximium M–P C. pubescens A, M untreated seeds (in A hilum terminal, M hilum medial) B treated seed C, G, K, O testa pattern of treated seeds showing anticlinal cell walls with fringe (C) and fibrils (G) D, H, L, M detail of testa cells E, I, N seeds with testa partly digested (in I hilum subterminal) F, J detail of a non-digested portion of the seed coat. Scale bars: 200 μm (A, B, E, I, M, N); 20 μm (C, D, G, H, J, K, L, P); 50 μm (F, O).

The distal end of the anticlinal cell walls may have three different types of appendages: (1) a thin ridge (Figs 9K, 10D, H, O, 11K, L, O, P, T, 12P, T, 14D, 15K), which is very common in the genus; (2) a more or less wide fringe (Figs 16K, L, R, S, 17O, Q, R, 18C); and (3) strands of thickenings differentiated as finger-like laciniations or fibrils (hair-like structures, Figs 9G, 10C, K, 14G, H, K, L, P, 17J, 18G, K).

The embryo in Capsicum is usually imbricate, meaning that the cotyledon tips are parallel or almost parallel to the radicle (Gunn and Gaffney 1974) and, less frequently, annular (cotyledon tips point toward the radicle, for example, C. dimorphum) or coiled (cotyledons coiled twice, Fig. 8B; for example, C. chacoense). The endosperm is firm, whitish and relatively abundant (Fig. 8B, C). Germination in Capsicum is epigeal.

To analyse the trichomes, temporary preparations of the epidermis of leaves, stems, calyx and corolla were made by making direct peels of the epidermis or cross sections; observations were made under light microscope and drawings were done with the help of a camara lucida.

Mature fruits were used to analyse the anatomy of the pericarp. Fresh fruits were collected in the wild, bought at various markets (domesticated species) or obtained from plants cultivated at the University of Cordoba (Argentina) (see Suppl. material 1: Appendix 1). Fruits were fixed in formalin-acetic acid-ethanol (FAA; 3.7% formaldehyde; 5% glacial acetic acid; 50% ethanol) for at least 48 hours and then small pieces of 3–5 × 3–5 mm from the middle part of the fruit were examined. For light microscopy, fixed material was dehydrated through an ascendant ethanol series (50°, 60°, 70°, 80°, 90°, 95°, 100°), clarified in pure xylene and embedded in Paramat^(TM). The samples were sectioned at 12 μm thick with a Minot rotary microtome. Sections were double-stained using Astra Blue/Basic Fuchsin (Kraus et. al. 1998; Zarlavsky 2014). Cross-sections were also made by hand. These were stained with safranin that stains intensely lignified cell walls (sclereids), which can then be easily differentiated from the cellulose-walled parenchyma cells. The cuticle was detected with Sudan IV. Outer and inner epidermal peels were obtained from the upper, middle and lower part of the fruit and were stained with safranin. The stained sections and peels were mounted in a drop of 50% glycerine solution. Measurements of the thickness of the cuticle and cuticular wedges, as well as the giant cells’ size, were made using a stage micrometer. Camera lucida drawings of pericarp sections are shown in many of the figures. Only dried fruits from herbarium specimens could be analysed for some species. Here, fruits were hydrated and kept in water at 90 °C for the necessary time (1–2 hours) to soften the tissues to be able to perform sectional cuts or pericarp peels.

Mature seed samples were taken from herbarium material or collected from wild or cultivated sources (see Suppl. material 2: Appendix 2). All seeds were selected and analysed in three ways for: (1) stereomicroscope (SM), (2) scanning electron microscopy (SEM) and (3) light microscope (LM) observations. The processing of the seeds for each case was as follows: (1) seeds were removed from fresh or hydrated fruits (untreated seeds) and gross morphology observations were made on dried seeds with a Zeiss Stemi 2000-C stereomicroscope (SM) at 32× magnification; (2) for scanning electron microscopy (SEM), seeds were prepared following the enzyme etching technique (Lester and Durrands 1984) to dissolve the outer cell walls, affixed to aluminium stubs with double-sided adhesive tape, then coated with gold and examined using either a JEOL JSM 35 CF SEM (LABMEM, National University of San Luis, Argentina) or a FE-SEM Sigma (LAMARX, National University of Córdoba, Argentina; (3) for sectioning for light microscopy, seeds were soaked in boiling water, washed with a diluted sodium hypochlorite solution (10%) to remove possible fungal hyphae and rinsed in distilled water; cross sections were free-hand cut with glass knives and then stained with Astra Blue and safranin (Johansen 1940). Terminology for seed (and embryo) morphology largely follows Gunn and Gaffney (1974), Zhang et al. (2005) and Chiou and Hastorf (2014).

Chromosome counts, cytogenetic information and DNA content are based on voucher specimens for which we were able to verify their correct identifications (vouchers cited in Table 2) or based on studies with taxonomic identity, verifiable from the descriptions material (vouchers not cited in Table 2). We do not cite all the extant cytogenetic literature in Table 2 for the domesticated species and some related wild species, since these works are not comparable with the karyotyping nomenclature systems (e.g. Levan et al. 1964; Moscone et al. 1996a) currently used; this means that some works have been excluded (Huskins and La-Cour 1930; Dixit 1931; Yamamoto and Sakai 1932; Tokunaga 1934; Raghavan and Venkatasubban 1940; Chennaveeraiah 1947; Sinha 1950; Vazart 1950, 1951; Eshbaugh 1964; Chennaveeraiah and Habib 1966; Shopova 1966; Datta 1968; Carluccio and Saccardo 1977; Limaye and Patil 1989; Alcorcés de Guerra 2001; Nascimento Sousa et al. 2015).

The monograph is based on results from many years of herbarium study and field work to collect these taxa across South America. Fresh material was preserved in FAA (formaldehyde–acetic acid–ethanol) or ethanol (70°) to perform measurements of reproductive organs using a Zeiss Stemi 2000-C stereomicroscope at 6.5–50× magnification or trichomes using a Leitz light microscope at 10–40× magnification. Descriptions were based on living plants observed during fieldwork and examination of ca. 6,900 herbarium specimens loaned from or inspected at 213 herbaria (acronyms follow Thiers 2021): A, AAU, AC, ALCB, ANDES, AS, ASE, ASSAM, B, BA, BAA, BAB, BAF, BHCB, BHZB, BKL, BM, BOLV, BR, C, CAL, CAS, CDS, CEN, CEPEC, CESJ, CHEP, CM, CNMT, COAH, COL, COLO, CONC, CONN, CORD, CR, CRI, CTES, CUVC, CUZ, DAV, DUKE, E, EAC, EAP, EFC, ENCB, ESA, ESAL, F, FCQ, FLAS, FLOR, FMB, FPS, FSU, FUEL, FURB, G, GA, GB, GBH, GH, GL, GOET, GUA, GUAY, HAL, HAJB, HAMAB, HAO, HAS, HB, HBG, HBR, HCF, HEH, HEPH, HFSL, HJ, HOXA, HRB, HRCB, HSB, HST, HSTM, HUA, HUAZ, HUCP, HUCS, HUEFS, HUEM, HUFU, HUFSJ, HULE, HURB, HURB, HUSA, HUSU, HUT, HVC, IAC, IAN, IEB, IBN, ICN, IND, INPA, IPA, ITIC, JAU, JAUM, JPB, K, KFTA, L, LAGU, LE, LINN, LP, LPB, LIL, LOJA, LUSC, M, MA, MAC, MCNS, MBM, MBML, MEDEL, MEL, MERL, MEXU, MICH, MISS, MNES, MO, MOL, MPU, MU, MY, NA, NDG, NY, OUPR, OXF, P, PACA, PEL, PEUFR, PH, PMSP, PR, PSO, PUL, PY, PYO, Q, QAP, QCA, QCNE, R, RB, REG, RIOC, RSA, RUSU, S, SAMES, SEV-H, SF, SMDB, SI, SJRP, SP, SPF, SPSF, TEFH, TUB, TULV, U, UB, UC, UDBC, UEC, UFG, UFP, UFRN, UNA, UNAH, UNOP, UNR, UOJ, UPCB, UPS, US, USCG, USF, USM, USZ, UT, VEN, VIC, VIES, VT, W, WAG, WU, WIS, XAL, YU and Z. Digital images were also accessed via different repositories, such as Global Plants (https://plants.jstor.org/), INCT Herbário Virtual (http://inct.splink.org.br), Herbário Virtual Reflora (http://reflora.jbrj.gov.br/reflora/herbarioVirtual), Atlas of the Florida Plants (http://florida.plantatlas.usf.edu) or Botanical Collections Databases of the following herbaria: B (http://ww2.bgbm.org/herbarium/default.cfm), COL (http://www.biovirtual.unal.edu.co/es/colecciones/search/plants/), F (http://fieldmuseum.org/explore/department/botany/collections), G (http://www.ville-ge.ch/cjb/bd.php), GH (https://huh.harvard.edu/pages/digital-resources), MA (http://colecciones.rjb.csic.es/), MPU (https://collections.umontpellier.fr/collections/botanique/herbier-mpu/base-herbier-mpu), NY, P (https://science.mnhn.fr/institution/mnhn/collection/p/item/search/form), S (https://herbarium.nrm.se/), US (https://collections.nmnh.si.edu/search/botany/), W (http://herbarium.univie.ac.at/database/search.php) and Z (http://www.herbarien.uzh.ch/index.html).

Measurements of dried material were made from dissections of flowers or fruits rehydrated in hot water, supplemented by measurements from living materials. Information about flower, fruit and seed colour was taken mainly from our own field observations and, in a few cases, flower colour was described from herbarium label data (e.g. C. hookerianum). The terminology used in the mature fruit descriptions of the domesticated species is based on the list of descriptors for Capsicum (IPGRI et al. 1995); pungency of immature and mature fruits was tested by tasting them in the field.

Distribution maps were produced using QGIS 3.16.0-Hannover (QGIS Development Team 2019) and were based on georeferenced data of all the herbarium collections analysed. Maps with boundaries of countries were imported from Natural Earth (www.naturalearthdata.com); elevation maps were obtained from the DIVA-GIS (http://diva-gis.org/gdata) spatial data. Specimens with latitude and longitude data on the labels were mapped directly; collections without geographical coordinates were georeferenced using different georeferencing tools, such as the point-radius method (determination of latitude and longitude and the maximum error distance), search for localities in Google Maps, Google Earth and GraphHopper Maps or in available databases with localities already georeferenced.

Preliminary conservation status was assessed using IUCN (2022) criteria B (for the relatively widespread species) and C (for some species with very restricted occurrences). We have given more weight to the extent of occurrence (EOO) in the threat assessments than to the area of occupancy (AOO), since the EOO is very sensitive to georeferencing bias and collecting effort (Knapp et al. 2020). The extent of occurrence and area of occupancy were calculated using the Geospatial Conservation Assessment Tool GeoCAT (Bachman et al. 2011; GeoCAT 2020).

Typification of cultivated taxa has been a particularly difficult task. For many taxa, the authors did not cite specimens or locality of the type. We searched for original material in potential herbaria and when we succeeded or duplicates were found, we designated lectotypes. In other taxa, lectotypifications were based on an illustration cited by the author in the protologue (e.g. Fingerhuth’s illustrations). For taxa recognised only as synonyms, we have cited the taxa in synonymy and indicated that duplicates have not been found rather than neotypifying these taxa. In cases where taxa were described from collections of living material cultivated in botanical gardens from unknown origin or the original material was destroyed in World War II, we designated neotypes when probable original material could be found (e.g. in C. ovatum) or using a modern collection (C. flexuosum). However, for the species described by Philip Miller (1768) in his“ Gardener’s Dictionary”, we have postponed typifications. Specimens made from plants grown by Miller at Chelsea Physic Garden in London (UK) are found in several different herbaria, mostly at BM and its associated historical herbaria; since we were unable to visit these herbaria for searching Miller’s original material due to restrictions for the current pandemic caused by Covid-19, we do not typify them here, leaving that for a separate study when these materials, including any non-digitised specimens, can be studied in detail.

In cases where specific herbaria have not been cited in protologues, we designate lectotypes rather than assuming holotypes exist (McNeill 2014). For cases in which the collection type was based on syntypes, we have examined as many as possible, choosing the best-preserved specimen as the lectotype; in all cases, we give the reasons for our choice in the species discussions.

Type specimens are cited with their barcodes in square brackets after each herbarium acronym, according to the style used in each herbarium (e.g. GOET003420 or MO-562486); we also indicate the sheet number after the barcode when available (i.e. IND-0153285, acc. # 139721). When barcodes are missing, we indicate only the sheet number (i.e. LIL acc. # 173409). In a few cases, we do not cite barcode or accession number (e.g. some type material from LE).

Identities of all numbered collections seen are presented in the List of Exsiccatae. Numbered and un-numbered collections are presented in Suppl. material 4: Appendix 4 where full citations of all the specimens examined for this treatment are presented in PDF format. We have not cited specimens of any Capsicum collected outside the Americas, with the exception of types.

Common names were taken from herbarium label data and reliable literature if we could verify the identity of taxa, but the list of common names for the cultivars of C. annuum, C. frutescens and C. chinense is not complete since we did not comprehensively examine the vast amount of literature where this information appears. Indigenous names are given in a separate paragraph for clarity indicating in brackets the indigenous language (if given). We cite only one specimen by provenance of the common and indigenous names per administrative division of each country. Uses as foods, spices or in ritual practices are cited in the species treatment and folk medicinal uses are summarised in Table 3, where the organ used and the medicinal properties, as well as the source of information, are indicated (verified specimens or literature).

All species are illustrated with line drawings, colour illustrations or both; photos were taken by the authors of this treatment or were provided by other colleagues (credits are cited in each case). For some species (C. caatingae, C. friburgense, C. hunzikerianum, C. mirum), photos were provided and taken in the field by members of the Associazione PepperFriends (Verona, Italy); these photos have no herbarium voucher, but the identification was verified by the senior author of this treatment (GEB).

Ecoregions were determined according to Olson et al. (2001). To determine the ecoregions and biomes occupied by the different species, maps were generated using QGIS 3.16.0-Hannover (QGIS Development Team 2019) superimposing layers of georeferenced data (as for the distribution maps, see above) with a layer of ecoregions obtained from Ecoregions2017^©Resolve (https://ecoregions.appspot.com/).