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2023 Volume 3
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ARTICLE   Open Access    

Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay

  • 1.

    Ondokuz Mayıs University, Faculty of Veterinary Medicine, 55200, Samsun, Türkiye

  • 2.

    Turkish Energy, Nuclear and Mineral Research Agency, Nuclear Energy Research Institute, 06980, Ankara, Türkiye

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  • The objective of the present study was to estimate applied doses of irradiated anchovy (1.83−4.22 kGy) and bluefish (1.98−5.40 kGy) for shelf-life extension by using image analysis in combination with comet assay for trading irradiated fish, especially for public health concerns. Comet assay was carried out considering the EN 13784 protocol. Observed comets were evaluated and recorded by the use of image analysis software (BS 200 ProP). Comets with long tails were observed in the irradiated anchovy and bluefish tissues. However, limited numbers of comets in intact cells were determined in the unirradiated samples. The obtained results clearly indicated that head diameter, head DNA, tail DNA, tail length, and tail moment parameters showed very good correlation (R2 = 0.82 and 0.77 for anchovy and bluefish respectively) with the applied doses of irradiation. Using image analysis in combination with comet assay may be a very useful tool for quantification of absorbed doses for local market checks and also in imported irradiated fish for official control at customs.

  • Introduction

    Scale insects (Hemiptera: Coccomorpha) with hypogeic habits are considered of high phytosanitary relevance for coffee crops (Rubiaceae: Coffea spp.) in Colombia[1]. A total of 65 species of scale insects associated with coffee roots have been recorded in Colombia[2−4]. The most species-rich family is the Pseudococcidae with 28 species distributed in nine genera: Dysmicoccus Ferris, 1950 (13 spp.), followed by Pseudococcus Westwood, 1840 (four spp.), Phenacoccus Cockerell, 1893 (three spp.), Planococcus Ferris, 1950, and Spilococcus Ferris, 1950 (two spp. each), and Chorizococcus McKenzie, 1960, Distichlicoccus Ferris, 1950, Ferrisia Fullaway, 1923, and Paraputo Laing, 1929 (one sp. each). For the family Rhizoecidae, 19 species have been recorded in six genera, namely, Rhizoecus Kunckel d'Herculais, 1878 (13 spp.), Pseudorhizoecus Green, 1933 (two spp.), and Capitisetella Hambleton, 1977, Coccidella Hambleton, 1946, Geococcus Green, 1902, and Ripersiella Tinsley, 1899 (one sp. each). Other minor families include Coccidae and Ortheziidae (five spp. in each family), Xenococcidae (three spp.), and Putoidae and Diaspididae (two spp. in each family) and Margarodidae (one sp.). For this study all previous records were re-analysed with the purpose of providing an accurate list of species

    The taxonomic identification of scale insects by a morphological approach is particularly difficult, mainly for two reasons. First, they are small insects (usually < 5 mm) that require the preparation of slide-mounted specimens. Second, the taxonomic keys needed for morphological identifications are primarily designed for adult female specimens[5]. Differing from other insect orders (e.g., Coleoptera, Diptera and Hymenoptera), female scale insects lack well-defined tagmata, as well as sclerites, sutures, or discernible areas. Characters of taxonomic value in scale insects include cuticular processes, such as pores, ducts and, setae[5]. Recognizing these cuticular structures on such small bodies poses a difficult task for non-expert entomologists. To facilitate accessible identification, this manuscript offers an illustrated taxonomic key to scale insect species associated with coffee roots in Colombia and is aimed at users with basic knowledge of scale insect morphology.

    Materials and methods

    A careful revision of the specimens studied by Caballero et al.[2], preserved in the Scale Insect Collection at the Entomological Museum 'Universidad Nacional Agronomía Bogota' UNAB (Bogotá, Colombia), was carried out to exclude species that are doubtfully recorded from coffee roots in Colombia. This re-assessment allowed the compilation of an accurate list of species that could be included in the taxonomic key. Additional species and information from Caballero[3] and Caballero et al.[4] also were used for construction of the key. List of species recorded for Colombia and c ollection data of specimens analized are in Supplemmental Table S1 and S2 respectivately.

    The illustrated taxonomic key (Table 1) is based on the external morphology of the adult female with a dichotomous structure. Each couplet after the first one is numbered followed by the number of the preceding couplet in parenthesis, e.g. 12(7) means that couplet 12 is derived from couplet 7; the numbers at the end of the couplet indicate the next couplet in order to arrive at the species name that best matches the character states selected by the user. It is illustrated in most of the steps using microphotographs. Acquisition and analysis of images were done with a Lumenera 1-5C camera and the software Image Pro Insight 8.0. Designs were performed with Affinity Photo V 2.1 and Affinity Designer V 2.1 software. The taxonomic keys were structured with some adaptations of published taxonomic keys[6−15]. The general morphological terminology follows Kondo & Watson[5] with specific terminology for Coccidae[6,16], Margarodidae[17], Ortheziidae[7], Diaspididae[18], Pseudococcidae, Putoidae[8,9], and Rhizoecidae[10,11]. The abdominal segmentation is given as SabdI for abdominal segment 1 to SabdVIII for abdominal segment 8. All microphotographs are of adult female scale insects or their taxonomically important morphological structures.

    Table 1.  Illustrated taxonomic key.
    No.DetailsRef.
    1Abdominal spiracles present (Fig. 1a)2
    −Abdominal spiracles absent (Fig. 1b)7
    Fig. 1 Abdominal spiracles (sp) on margin (a) present on Eurhizococcus colombianus, (b) absent on Distichlicoccus takumasai.
    2(1)Anal aperture without pores and setae (Fig. 2a); legs shorter than half of the transversal diameter of body (Fig. 2b); eyespots and mouthparts absentEurhizococcus colombianus

    Jakubski, 1965
    −Anal aperture forming a well-developed anal ring with pores and setae (Fig. 2c); legs longer than transversal diameter of body; eyespots and mouthparts present (Fig. 2d)
    3
    Fig. 2 Eurhizococcus colombianus: (a) Anal aperture without pores and setae in the border, (b) section of mid body showing the length of hind leg (lel) and transversal body line (btl). Insignorthezia insignis: (c) Anal aperture with pores (po) and setae (st), (d) section of head with protruding eyespot (es) and labium (lb).
    3(2)Antennae each with eight segments (Fig. 3a)4
    −Antennae each with fewer than five segments (Fig. 3b)5
    Fig. 3 (a) Eight-segmented antenna. (b) Four-segmented antenna.
    4(3)Transversal bands of spines absent in ventral region surrounded by an ovisac band (Fig. 4a); dorsal interantennal area without sclerosis (Fig. 4b)Insignorthezia insignis (Browne, 1887)
    −Transversal bands of spine plates present in ventral region surrounded by an ovisac band (Fig. 4c); longitudinal sclerosis on dorsum in interantennal area (Fig. 4d)Praelongorthezia praelonga (Douglas, 1891)
    Fig. 4 Insignorthezia insignis: (a) Abdomen without transversal clusters of wax plates, (b) Dorsal interantennal area without sclerosis. Praelongorthezia praelonga: (c) Abdomen with transversal clusters of wax plates marked by dash lines, (d) dorsal interantennal area with a longitudinal sclerotic plate (ep).
    5(3)Antennae each with three segments (Fig. 5a)Newsteadia andreae Caballero, 2021
    −Antennae each with four segments (Fig. 5b)6
    Fig. 5 (a) Three-segmented antenna of Newsteadia andreae. Note the presence of pseudosegmentation which gives the appearance of additional segments in the last antennal segment. (b) Four-segmented antenna of Mixorthezia minima.
    6(5)Dorsal area anterior to anal ring with simple pores on protuberances (Fig. 6a); ventral areas surrounding each coxa with a row of wax plate spines (Fig. 6b)Mixorthezia minima Koczné Benedicty & Kozár, 2004
    −Dorsal area anterior to anal ring without simple pores or protuberances (Fig. 6c); ventral areas posterior to each coxa without wax plate spines (Fig. 6d)Mixorthezia neotropicalis (Silvestri, 1924)
    Fig. 6 Mixorthezia minima: (a) Dorsum of area anterior to anal ring with close-up of simple pores on protuberances (dash box); (b) ventral area posterior to each coxa with a row of wax plate spines (dash box). Mixorthezia neotropicalis: (c) Close-up of dorsum of area anterior to anal ring lacking simple pores on protuberances (dash box); (d) ventral area posterior to each coxa without associated wax plate spines.
    7(1)Anal plates present (Fig. 7a)8
    −Anal plates absent (Fig. 7b)12
    Fig. 7 (a) Anal apparatus of Saissetia coffeae with anal plates (ap) covering the anal aperture (aa). (b) Anal apparatus of Pseudococcus sp. with anal aperture lacking anal plates.
    8(7)Antennae and legs with length similar to or shorter than spiracles (Fig. 8a)9
    −Antennae and legs with length at least twice as long as spiracles (Fig. 8b)11
    Fig. 8 (a) Antenna (an) and foreleg (lg) (green lines), and anterior spiracle (sp) (yellow line) of Toumeyella coffeae showing their relative length. Note the similar size of the limbs and spiracle. (b) Antenna (an) and leg (lg) (green lines), and anterior spiracle (sp) (yellow line) of Coccus viridis showing their relative length. Note the relatively smaller size of the spiracle.
    9(8)Ventral tubular macroducts present (Fig. 9)Toumeyella coffeae
    Kondo, 2013
    −Ventral tubular macroducts absent10
    Fig. 9 Ventral tubular macroducts (dash box) and close-up of macroducts (photo on right side).
    10(9)Orbicular pores (Fig. 10a) and cribriform platelets present (Fig. 10b); dorsal setae absent; opercular pores absentCryptostigma urichi (Cockerell, 1894)
    −Orbicular pores and cribriform platelets absent; dorsal setae present (Fig. 10c); numerous opercular pores present throughout mid areas of dorsum (Fig. 10d)Akermes colombiensis Kondo & Williams, 2004
    Fig. 10 Cryptostigma urichi: (a) Orbicular pore and (b) close-up of a cribriform platelet. Akermes colombiensis: (c) Close-up of a dorsal body setae (dash box) and (d) close-up of opercular pores (arrows).
    11(8)Band of ventral tubular ducts in lateral and submarginal regions absent, ventral tubular ducts of one type; anal plates without discal setae (Fig. 11a); dorsal body setae capitate or clavate (Fig. 11b); perivulvar pores with seven or eight loculi, rarely with 10 loculi (Fig. 11c)Coccus viridis
    (Green, 1889)
    −Band of ventral tubular ducts in lateral and submarginal regions present, submarginal region with two types of tubular ducts (Fig. 11d); anal plates with discal setae (Fig. 11e); dorsal body setae spine-like, apically pointed (Fig. 11f); perivulvar pores mostly with 10 loculi (Fig. 11g)Saissetia coffeae
    (Walker, 1852)
    Fig. 11 Coccus viridis: (a) Anal plates without discal setae; (b) dorsal body setae capitate (top) or clavate (below); (c) multilocular disc pores mostly with eight loculi. Saissetia coffeae: (d) Ventral submarginal region with two types of tubular ducts; (e) each anal plate with a discal seta; (f) dorsal body setae acute; (g) multilocular disc pores with mostly 10 loculi.
    12(7)Cerarii present on body margin, at least a pair on each anal lobe (Fig. 12a)13
    −Cerarii absent on body margin (Fig. 12b)38
    Fig. 12 Abdominal body margin of (a) Pseudococcus sp. with three cerarii (dash box) and (b) Rhizoecus sp. (dash box) without cerarii.
    13(12)Enlarged oral collar tubular ducts composed of a sclerotized area surrounding the border and a set of flagellated setae (Ferrisia-type oral collar tubular ducts) (Fig. 13a)Ferrisia uzinuri
    Kaydan & Gullan, 2012
    −Oral collar tubular ducts simple, not as above (Fig. 13b) or absent14
    Fig. 13 (a) Ferrisia-type oral collar tubular ducts with aperture of tubular duct (ad) surrounded by a sclerotized area (sa) and associated flagellate setae (fs). (b) Oral collar tubular ducts simple (arrows).
    14(12)Antenna with nine segments (Fig. 14a)15
    −Antenna with eight segments (Fig. 14b) or fewer (Fig. 14c)19
    Fig. 14 Antenna with (a) nine segments, (b) eight segments and (c) seven segments.
    15(14)Cerarii with more than five conical setae (Fig. 15a); hind trochanter with six sensilla, three on each surface (Fig. 15b)16
    −Cerarii with two lanceolate setae (Fig. 15c); hind trochanter with four sensilla, two on each surface (Fig. 15d)17
    Fig. 15 Puto barberi: (a) upper and lateral view of a cerarius, (b) close-up of the surface of trochanter with three sensilla (arrows). Phenacoccus sisalanus: (c) cerarius, (d) trochanter with two sensilla (arrows) on single surface.
    16(15)Cerarii with tubular ducts (Fig. 16a)Puto antioquensis
    (Murillo, 1931)
    −Cerarii without tubular ducts (Fig. 16b)Puto barberi
    (Cockerell, 1895)
    Fig. 16 (a) Cerarius associated with tubular ducts (arrows). (b) Cerarius without tubular ducts.
    17(15)Oral collar tubular ducts absentPhenacoccus sisalanus Granara de Willink, 2007
    −Oral collar tubular ducts present, at least on venter (Fig. 17)18
    Fig. 17 Ventral surface with oral collar tubular ducts (dash circles).
    18(17)Oral collar tubular ducts restricted to venterPhenacoccus solani
    Ferris, 1918
    −Oral collar tubular ducts present on dorsum and venterPhenacoccus parvus Morrison, 1924
    19(14)Oral rim tubular ducts present (Fig. 18)20
    −Oral rim tubular ducts absent26
    Fig. 18 Oral rim tubular ducts in upper view (dash circles) and close-up of lateral view.
    20(19)Oral rim tubular ducts present on venter onlyPseudococcus landoi (Balachowsky, 1959)
    −Oral rim tubular ducts present on both dorsum and venter21
    21(20)Cerarii restricted to anal lobes (Fig. 19a)Chorizococcus caribaeus Williams & Granara de Willink, 1992
    −Cerarii present, at least on the last five abdominal segments (Fig. 19b)22
    Fig. 19 Location of cerarii (dash boxes) on abdominal margin with close-up of cerarius (a) restricted to anal lobes (dash boxes) and (b) cerarii present on the last five abdominal segments.
    22(21)Circulus absent (Fig. 20a)23
    −Circulus present (Fig. 20b)24
    Fig. 20 Ventral mid area of abdominal segments III and IV (dash box) of (a) Distichlicoccus takumasai without circulus and (b) Pseudococcus jackbeardsleyi with circulus.
    23(22)Multilocular disc pores present on venter of SabdIV and posterior segments (Fig. 21a); hind coxa with translucent pores and hind femur without translucent pores (Fig. 21b)Spilococcus pressus
    Ferris, 1950
    −Multilocular disc pores absent, if some present, not more than three around vulvar opening (i.e. venter of SabdVII or SabdVIII); hind coxa without translucent pores (Fig. 21c) and hind femur with translucent pores (Fig. 21d)Distichlicoccus takumasai Caballero, 2021
    Fig. 21 Spilococcus pressus: (a) Ventral section of abdomen with multilocular disc pores (arrows); (b) hind leg with close-up of coxa with translucent pores (arrows). Distichlicoccus takumasai: (c) Hind coxa without translucent pores; (d) hind femur with translucent pores (arrows).
    24(22)Eyes without discoidal pores nor sclerotized surrounding area (Fig. 22a); circulus with transversal diameter 40 to
    60 µm (Fig. 22b)    		Pseudococcus luciae Caballero, 2021
    −Eyes with discoidal pores and sclerotized surrounding area (Fig. 22c); circulus diameter 100 to 200 µm (Fig. 22d)26
    25(24)Oral rim tubular ducts on dorsal abdominal segments numbering three to eight; area between posterior ostiole and cerarius of SabdVII without oral rim tubular ducts (Fig. 23a)Pseudococcus elisae Borchsenius, 1947
    −Oral rim tubular ducts on dorsal abdominal segments numbering 14 to 27; area between posterior ostiole and cerarius of SabdVII with an oral rim tubular duct (Fig. 23b)Pseudococcus jackbeardsleyi Gimpel & Miller, 1996
    Fig. 22 Pseudococcus luciae: (a) Eyespot without surrounding sclerotized area nor associated pores; (b) circulus ca. 58 µm wide. Pseudococcus jackbeardsleyi: (a) Eyespot with sclerotized area (sa) and associated pores (po); (d) circulus ca. 154 µm wide.
    Fig. 23 (a) Dorsal margin of abdominal segments VI to VIII, between cerarius of anal lobe (C1), cerarius of SabdVII (C2) and posterior ostiole (os) without oral rim tubular ducts. (b) Dorsal margin of abdominal segments VI to VIII, between cerarius of anal lobe (C1), cerarius of SabdVII (C2) and posterior ostiole (os) with an oral rim tubular duct and/or cerarius adjacent to SabdVII.
    26(19)Oral collar tubular ducts (Fig. 24) on both dorsum and venter27
    −Oral collar tubular ducts restricted to venter28
    Fig. 24 Oral collar tubular duct in lateral view.
    27(26)Hind coxa with translucent pores (Fig. 25a); anal lobe with sclerotized bar, not on a sclerotized area (Fig. 25b); multilocular disc pores present posterior to fore coxaPlanococcus citri-minor complex
    −Hind coxa without translucent pores (Fig. 25c); anal lobe without sclerotized bar, on a sclerotized area (Fig. 25d); multilocular disc pores absent posterior to fore coxaDysmicoccus quercicolus (Ferris, 1918)
    28(27)Oral collar tubular ducts absent on venter of both head and thorax.29
    −Oral collar tubular ducts present on either head or thorax, but not on both areas (Fig. 26)30
    Fig. 25 Planococcus citri-minor complex: (a) Hind coxa with translucent pores (dash box) and (b) anal lobe with a sclerotization forming a bar (ab). Dysmicoccus quercicolus: (c) Hind coxa without translucent pores and (d) anal lobe with irregular broad sclerotized area (sa).
    Fig. 26 Marginal area of Dysmicoccus grassii, lateral to posterior spiracle (ps), with close-up of oral collar tubular ducts (oc) (left side).
    29(28)Translucent pores present on hind coxa, trochanter, femur and tibia (Fig. 27a); marginal clusters of oral collar tubular ducts on venter of SabdVI and SabdVIIDysmicoccus caribensis Granara de Willink, 2009
    −Translucent pores restricted to hind femur and tibia (Fig. 27b); marginal clusters of oral collar tubular ducts present on venter of SabdIV to SabdVIIParaputo nasai
    Caballero, 2021
    Fig. 27 (a) Hind leg of Dysmicoccus caribensis with translucent pores on coxa (cx), trochanter (tr) and femur (fm), and tibia (tb). (b) Hind leg of Paraputo nasai with translucent pores restricted to femur (fm) and tibia (tb).
    30(28)Hind coxa with translucent pores (Fig. 28a)Dysmicoccus sylvarum
    Williams & Granara de Willink, 1992
    −Hind coxa without translucent pores (Fig. 28b)31
    Fig. 28 (a) Translucent pores on hind coxa. (b) Translucent pores absent on hind coxa.
    31(30)Hind trochanter with translucent pores (Fig. 29a)Dysmicoccus varius
    Granara de Willink, 2009
    −Hind trochanter without translucent pores (Fig. 29b)32
    Fig. 29 Translucent pores (a) on hind trochanter, (b) absent from hind trochanter.
    32(31)Oral collar tubular ducts present on margin of thorax (Fig. 30)33
    −Oral collar tubular ducts absent from margin of thorax34
    Fig. 30 Prothorax margin of Dysmicoccus grassii with close-up of oral collar tubular ducts.
    33(32)Multilocular disc pores absent on SabdV; dorsal area immediately anterior to anal ring with tuft of flagellate setae; longest flagellate seta as long as diameter of anal ring (Fig. 31a), and discoidal pores larger than trilocular pores (Fig. 31b)Dysmicoccus radicis
    (Green, 1933)
    −Multilocular disc pores present on SabdV; dorsal area immediately anterior to anal ring without a tuft of flagellate setae; flagellate setae much shorter than diameter of anal ring (Fig. 31c) and discoidal pores smaller than trilocular pores (Fig. 31d)Dysmicoccus grassii (Leonardi, 1913)
    34(32)Oral collar tubular ducts absent in interantennal area35
    −Oral collar tubular ducts present in interantennal area (Fig. 32)36
    35(34)Translucent pores on hind leg restricted to tibia (Fig. 33a)Dysmicoccus perotensis
    Granara de Willink, 2009
    −Translucent pores on hind leg present on tibia and femur (Fig. 33b)Dysmicoccus joannesiae-neobrevipes complex
    Fig. 31 Dysmicoccus radicis: (a) Area anterior to anal ring with a cluster of flagellate setae (fs) and anal ring (ar) showing the diameter of the different pores (dash box); (b) discoidal pores (dp) and trilocular pores (tp). Dysmicoccus grassii: (c) Area anterior to anal ring with scattered short flagellate setae (fs) contrasted with anal ring (ar) diameter (dash box); (d) discoidal pores (dp) and trilocular pores (tp) with similar diameter.
    Fig. 32 Interantennal area (dash box) of Dysmicoccus brevipes with close-up of oral collar tubular ducts.
    Fig. 33 (a) Hind leg of Dysmicoccus perotensis with close-up of femur and tibia with translucent pores on tibia only (arrows). (b) Hind leg of Dysmicoccus joannesiae-neobrevipes complex with close-up of femur and tibia with translucent pores (arrows).
    36(34)Hind coxa with translucent pores (see Fig. 28a)Dysmicoccus mackenziei
    Beardsleyi, 1965
    −Hind coxa without translucent pores (see Fig. 28b)37
    37(36)Dorsal SabdVIII setae forming a tuft-like group, each seta conspicuously longer than remaining dorsal abdominal setae (Fig. 34a) and setal length similar to anal ring diameter (60–80 µm long)Dysmicoccus brevipes (Cockerell, 1893)
    −Dorsal SabdVIII setae evenly distributed, each setae as long as remaining dorsal abdominal setae (Fig. 34b) and length less than half diameter of anal ringDysmicoccus texensis-neobrevipes complex
    38(12)Tritubular ducts absent39
    −Tritubular ducts present (Fig. 35a-b)46
    Fig. 34 (a) Abdomen of Dysmicoccus brevipes with dorsal setae on SabdVIII (lfs) longer than setae on anterior segments (sfs). (b) Abdomen of Dysmicoccus texensis-neobrevipes complex with dorsal setae (ufs) along the abdominal segments of uniform length and scattered distribution.
    Fig. 35 (a) Tritubular duct in upper (left) and lateral view (right) with the border of the cuticular ring attached to tubules. (b) Tritubular duct with the border of the cuticular ring widely separated from tubules (arrows).
    39(38)Anal lobes strongly protruded, bulbiform (Fig. 36a) jutting out from margin for a distance equivalent to diameter of anal ring40
    −Anal lobes shallow, if protruded, their length never more than half of diameter of anal ring (Fig. 36b)42
    Fig. 36 (a) Abdomen of Neochavesia caldasiae with anal lobes (al) protruding beyond the anal aperture (aa). (b) Abdomen of Ripersiella sp. with anal lobes (al) at the same level as the anal aperture (aa).
    40(39)Anal aperture located at the same level as the base of anal lobes (Fig. 37a); antennae located on ventral margin of headNeochavesia caldasiae (Balachowsky, 1957)
    −Anal aperture located anterior to bases of anal lobes (Fig. 37b); antennae located on dorsum of head41
    Fig. 37 (a) Abdomen of Neochavesia caldasiae with anal aperture (aa) positioned between the anal lobes (al), at the same level as the bases of anal lobes (dash line). (b) Abdomen of Neochavesia eversi with anal aperture (aa) situated anterior to the bases of the anal lobes (al) (dash line).
    41(40)Antennae each with five segments, situated on a membranous base (Fig. 38a); length of hind claw less than length of hind tarsus (Fig. 38b)Neochavesia trinidadensis (Beardsley, 1970)
    −Antennae each with four segments, situated on a sclerotized base (Fig. 38c); hind claw longer than hind tarsus (Fig. 38d)Neochavesia eversi (Beardsley, 1970)
    Fig. 38 (a) Antenna with four segments and a membranous base (mb). (b) Hind tarsus (green line) longer than the hind claw (red line). (c) Antenna with four segments and a sclerotized base (sb). (d) Hind tarsus (green line) shorter than hind claw (red line).
    42(39)Body setae capitate, at least on one surface (Fig. 39a)43
    −Body setae never capitate (Fig. 39b)44
    Fig. 39 (a) Capitate setae. (b) Flagellate setae.
    43(42)Anal aperture without associated cells (Fig. 40a); three-segmented antennae (Fig. 40b); ventral setae in median
    and submedian regions capitate    		Capitisitella migrans
    (Green, 1933)
    −Anal aperture surrounded by cells (Fig. 40c); six-segmented antennae (Fig. 40d); ventral setae in medial and submedial regions flagellateWilliamsrhizoecus coffeae
    Caballero & Ramos, 2018
    44(42)Three-segmented antennae (Fig. 41a); circulus present (Fig. 41b)Pseudorhizoecus bari
    Caballero & Ramos, 2018
    −Five-segmented antennae (Fig. 41c); circulus absent45
    Fig. 40 Capitisitella migrans: (a) Anal aperture of surrounded only by setae; (b) antenna composed of three segments. Williamsrhizoecus coffeae: (c) Anal aperture of surrounded by setae and cells (flesh); (d) antenna composed of six segments.
    Fig. 41 Pseudorhizoecus bari: (a) Antenna composed of three segments and (b) circulus. (c) Antenna of Pseudorhizoecus proximus composed of five segments.
    45(44)Multilocular disc pores absent; anal aperture ornamented with small protuberances and two to five short setae, each seta never longer than 1/3 diameter of anal aperture, without cells (Fig. 42a)Pseudorhizoecus proximus
    Green, 1933
    −Multilocular disc pores present (Fig. 42b); anal aperture not ornamented with small protruberances, ring with well-developed cells and six long setae, each seta as long as diameter of anal ring (Fig. 42c)Ripersiella andensis (Hambleton,
    1946)
    Fig. 42 (a) Anal aperture of Pseudorhizoecus proximus surrounded by protuberances (pr) and a few short setae (st). Ripersiella andensis: (b) Ventral section of abdomen with multilocular disc pores (mp); (c) anal aperture with a ring of cells and six long setae (se).
    46(38)Anal lobes strongly protruded, conical, each one with a stout spine at apex (Fig. 43a)Geococcus coffeae
    Green, 1933
    −Anal lobes flat or barely protruded, without spines at apex (Fig. 43b)47
    47(46)Venter of abdomen with clusters of trilocular pores in medial region (Fig. 44a)Coccidella ecuadorina Konczné Benedicty & Foldi, 2004
    −Venter of abdomen with trilocular pores evenly dispersed, never forming clusters in medial region (Fig. 44b)48
    Fig. 43 (a) Abdomen of Geococcus coffeae with protruding anal lobe (al) with a stout spine at the apex (sp). (b) Abdomen of Rhizoecus sp. with anal lobe (al) flat, with numerous flagellate setae (fs) at the apex.
    Fig. 44 (a) Ventral surface of Coccidella ecuadorina with clusters of trilocular pores (tc) (dash box) on medial region of abdomen. (b) Ventral surface of Rhizoecus sp. with trilocular pores (tr) scattered on venter of abdomen.
    48(47)Antennae with six well-developed segments (Fig. 45a)51
    −Antennae with five well-developed segments (Fig. 45b), apical segment sometimes partially divided (Fig. 45c)49
    Fig. 45 (a) Six-segmented antenna. (b) Five-segmented antenna. (c) Five-segmented antenna with partially divided apical segment (pd). Note: antennal segments numbered in Roman numerals.
    49(48)Antennae length more than 140 µm (Fig. 46a); tritubular ducts of similar diameter to trilocular pores (± 2 µm variation) (Fig. 46b); tritubular ducts with space between ductules and edge as wide as the ductules (Fig. 46c); slender ductule, width/length ratio 1:6Rhizoecus coffeae
    Laing, 1925
    −Antennae length less than 130 µm (Fig. 46d); tritubular ducts of diameter nearly twice diameter of trilocular pores (Fig. 46e); tritubular ducts with reduced space or without space between ductules and edge (Fig. 46f); stout ductule, width/length ratio 1:350
    50(49)Tubular ducts present (Fig. 47a); each anal lobe with around 28 dorsal setae of similar length, greater than 30 µm (Fig. 47b, al); and dorsal marginal clusters of setae on SabdVII 20–30 µm long (Fig. 47b, SabdVII)Rhizoecus setosus (Hambleton, 1946)
    −Tubular ducts absent; each anal lobe with around 14 dorsal setae, with length less than 15 µm (Fig. 47c, al); dorsal marginal clusters of setae on SabdVII with length less 15 µm (Fig. 47c, SabdVII)Rhizoecus compotor
    Williams & Granara de Willink, 1992
    Fig. 46 (a) Antenna ca. 207 µm long. (b) Tritubular ducts (td) and trilocular pores (tp) with similar diameter. (c) Close-up of a tritubular duct indicating the space between the cuticular ring (mg) and the ductule (dt). (d) Antenna ca. 105 µm long. (e) Each tritubular duct (td) twice the diameter of a trilocular pore (tp). (f) Close-up of tritubular duct without a space between the cuticular ring (mg) and the ductule (dt).
    Fig. 47 Rhizoecus setosus: (a) Tubular ducts (td); (b) anal lobe (al) and abdominal segment (SabdVII) with marginal clusters of setae longer than 30 µm. (c) Abdomen of Rhizoecus compotor with marginal cluster of setae shorter than 20 µm on anal lobe (al) and abdominal segment (SabdVII).
    51(48)Fore tibia with at least one of two internal preapical setae spine-like (Fig. 48a-b)52
    −Fore tibia with both internal preapical setae flagellate (Fig. 48c)56
    Fig. 48 Fore legs with preapical setae on tibia (ft): (a) one flagellate (fs) and one spine seta (ss), (b) with a pair of spine setae (ss), (c) with a pair of flagellate setae (fs).
    52(51)Fore tibia with one internal preapical spine-like setae and other seta flagellate (Fig. 48a); anal ring composed of spine-like setae (Fig. 49a); circulus absentRhizoecus spinipes (Hambleton, 1946)
    −Fore tibia with both internal preapical setae spine-like (Fig. 48b); anal ring composed of flagellate-like setae (Fig. 49b); at least, one circulus present (Fig. 49c)53
    Fig. 49 (a) Anal ring (ar) of Rhizoecus spinipes with spine-like setae (ss). (b) Anal ring (ar) of Rhizoecus arabicus with flagellate setae (fs). (c) Circulus of Rhizoecus cacticans.
    53(52)Claw digitules setose and short, length less than half length of claw (Fig. 50a)54
    −Claw digitules capitate and long, as long as claw (Fig. 50b)55
    Fig. 50 Claw with claw digitule: (a) setose (sd), (b) flagellate (fd).
    54(53)Anal ring with external row composed of 35 cells or more (Fig. 51a, ext); anal ring with external and internal rows separated by a space as wide as a cell of the external row (Fig. 51a, spc); anal ring cells without spicules (Fig. 51a, sp)Rhizoecus variabilis Hambleton, 1978
    −Anal ring with external row composed of less than 30 cells (Fig. 51b, ext); anal ring with external and internal rows separated by a narrow space, as wide as half (or less) a cell of the external row (Fig. 51b, spc); anal ring cells with spicules (Fig. 51b, sp)Rhizoecus arabicus Hambleton, 1976
    Fig. 51 (a) Anal ring of Rhizoecus variabilis with external row (ext) of anal ring consisting of over 35 cells; external row separated from the internal row (int) by a similar width as the diameter of a cell (spc). (b) Anal ring of Rhizoecus arabicus with external row (ext) of anal ring with less than 30 cells; external row separated from the internal row (int) by a width less than half the diameter of a cell (spc); cells of the external row with spicules (sp).
    55(53)More than 80 tritubular ducts; circulus with basal diameter at least five times greater than apical diameter (Fig. 52a); stick-like genital chamber, parallel borders and all of similar width and structure, length across about two abdominal segments (169–175 µm long) (Fig. 52b)Rhizoecus atlanticus (Hambleton, 1946)
    −Less than 50 tritubular ducts; circulus with basal diameter less than three times the apical diameter (Fig. 52c); genital chamber with basal third two times wider than anterior two-thirds, length across one abdominal segment (43–52 µm long) (Fig. 52d)Rhizoecus cacticans (Hambleton, 1946)
    Fig. 52 Rhizoecus atlanticus: (a) Circulus with diameter at base five times the apical diameter, (b) genital chamber tubular shape, length ca. 150 µm long. Rhizoecus cacticans: (c) Circulus with diameter at base about two times the apical diameter, (d) genital chamber with proximal section basiform and distal section tubular, with arms, length ca. 45 µm long.
    56(51)Multilocular disc pores absent on dorsumRhizoecus mayanus (Hambleton, 1946)
    −Multilocular disc pores present on dorsum57
    57(56)Marginal prothoracic setae length greater than 50 µm (Fig. 53a); marginal SabdVII setae length greater than 45 µm (Fig 53b)Rhizoecus colombiensis Ramos-Portilla & Caballero, 2016
    −Marginal prothoracic setae length less than 25 µm (Fig. 53c); marginal SabdVII setae length less than 30 µm (Fig. 53d)58
    Fig. 53 Rhizoecus colombiensis: (a) Body margin with a long seta (pts) (> 40 µm), longer than remaining setae in prothorax; (b) margin of abdominal segment VII (SabdVII) (st). with a long seta (pts) (> 40 µm), longer than remaining setae in abdomen. Rhizoecus americanus: (c) Margin of prothorax (pts) with setae of uniform length, shorter than 30 µm; (d) margin of abdominal segment VII (SabdVII) with setae (st) shorter than 30 µm.
    58(57)Tritubular ducts of two sizesRhizoecus caladii
    Green, 1933
    −Tritubular ducts of three sizesRhizoecus americanus (Hambleton, 1946)
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    Results

    The following illustrated taxonomic key (Table 1) is a tool for the identification of adult female scale insects (Hemiptera : Sternorrhyncha : Coccomorpha) associated with coffee roots in Colombia, which includes 59 species from seven families (see Supplemental Table S1).

    Discussion

    The taxonomic key includes 59 species associated with coffee roots. Hemiberlesia sp., Odonaspis sp., Rhizoecus stangei McKenzie, 1962, Spilococcus mamillariae (Bouche, 1844), Planococcus citri (Risso, 1813) and Planococcus minor (Maskell, 1897) were excluded from the key. In the case of the two armoured scale insects, the specimens were found in Berlese funnel samples associated with coffee roots[2], however, there is no evidence of these species feeding on the roots and there are no previous records of association of Hemiberlesia nor Odonaspis species with coffee roots.

    Previous records of single specimens of R. stangei and S. mamillariae by Caballero et al.[2] were determined as misidentifications of Rhizoecus caladii Green, 1933 and Spilococcus pressus Ferris, 1950, respectively. Spilococcus mamillariae is considered as an oligophagous species, but mainly associated with Cactaceae plants and feeding on the aerial parts of plants[19,20]. There are no records of S. mamillariae being found on any plant species of the family Rubiaceae, and hence we have removed this species from the list of species associated with coffee roots. The species R. stangei, which has been recorded only from Mexico and lacks information on its host plant[21] has apparently not been found since its original description[8].

    Planococcus citri and Pl. minor were listed also by Caballero et al.[2] as literature records. the morphological identification of P. citri and P. minor needs to be complemented with molecular and geographical analysis to be more accurate[22]. Therefore, the present key considers only identification to the Planococcus citri-minor complex.

    Furthermore, many specimens of Dysmicoccus collected from coffee roots in Colombia have morphological character states that overlap with Dysmicoccus neobrevipes Beardsley, 1959, Dysmicoccus joannesiae (Costa Lima, 1939) and Dysmicoccus texensis (Tinsley, 1900). The first case is a mix of character states of D. texensis and D. neobrevipes. The number of setae in the abdominal cerarii and the size of oral collar tubular ducts are the most important characters used to differentiate the adult females of Dysmicoccus species[8,23]. Adult females of D. texensis have a consistent pattern of only two setae in all thoracic and abdominal cerarii, along with a uniform size of oral collar tubular ducts (OC). On the other hand, D. neobrevipes varies in the number of setae in the cerarii, ranging from two to seven, accompanied by two distinct sizes of OC. These character states are generally constant among specimens found on the aerial parts of plants. However, among the specimens examined here, while the anal lobe cerarii consistently have two setae on the specimens of D. texensis found on the roots, the remaining cerarii display a variable number of setae, notably ranging from two to five, particularly within the abdominal cerarii. Furthermore, the OC of these specimens all are the same size. Regarding the differences in number of setae in the cerarii, Granara de Willink[23] underlined the need of more comprehensive studies to definitively separate these species.

    The second case involves D. joannesiae and D. neobrevipes. These species exhibit similarities in the number of setae on each cerarius (ranging from two to seven setae per cerarius) and differences in the number of clusters of OC along the abdominal margin; D. joannesiae has more than 25 clusters of OC and D. neobrevipes has fewer than 10 clusters of OC[8]. Granara de Willink also separated these two species by the presence of OC on the thorax and head[23] (present in D. neobrevipes and absent in D. joannesiae). Within the specimens of putative D. neobrevipes studied here, a few had clusters of OC numbering 15 to 20 along the abdominal margin and OC on the thorax and head. The primary challenge with addressing this dilemma lies in the fact that D. joannesiae has only been reported on Joannesia princeps Vell., 1798 (Euphorbiaceae) in Brazil and on Annona muricata (Annonaceae) intercepted in London from Saint Lucia[8,24]. Moreover, there has been no additional morphological variations recorded in the new records of D. joannesiae since its initial description in 1932 by Costa Lima. Therefore, the character states defining D. joannesiae are based on six type specimens. Based on these arguments, the following taxonomic key considers two species complex groups, namely the Dysmicoccus texensis-neobrevipes complex and the D. joannesiae-neobrevipes complex.

    Following article 31.1.2 of the International Commission of Nomenclature (ICZN), herein we make a change in nomenclature for Distichlicoccus takumasae Caballero, 2021. The ending -ae for takumasae is incorrect because the species was dedicated to Dr. Takumasa Kondo (a male coccidologist), and thus the correct ending is -i, hence the species epithet is herein amended to 'takumasai'. The corrected name is Distichlicoccus takumasai Caballero, 2021.

    Conclusions

    After reviewing the species of scale insects associated with coffee roots in Colombia, we have compiled a list of 59 species (Supplemental Table S1). Although this study did not focus on the effect of habit (aerial vs underground) or host plant on the morphology of scale insects, we detected significant morphological variation within facultative hypogeal species. Until further studies allow an understanding of the overlap of character states between D. texensis – D. neobrevipes and D. joannesiae – D. neobrevipes, we suggest considering these species as a morphological complex for hypogeal specimens. Further ecomorphological studies should be conducted to determine whether the morphology of a species may differ when feeding on the aerial parts compared when feeding on the underground parts of a host and to try to elucidate what factors trigger those changes, especially in species associated with coffee plants. As for the species complex, further collecting, morphological, and molecular studies should help elucidate these taxonomic problems.

    During the literature review performed for this study, we realized that most of the records of species are limited to mentioning the host but not the plant part on which collections were made, however, it is suspected that most species are normally collected from the aerial parts of the plant host. Although this taxonomic key is limited to root-associated species recorded in Colombia, this key could be useful for identifying scale insects associated with coffee in other tropical regions, extending also to species collected from the aerial parts of the hosts.

    Author contributions

    The authors confirm contributions to the paper as follows: study conception and design: Caballero A, Kondo T; data collection: Caballero A, Kondo T; analysis and interpretation of results: Caballero A, Kondo T; draft manuscript preparation: Caballero A, Kondo T. Both authors reviewed the results and approved the final version of the manuscript.

    Data availability

    The data (microscopy slides of specimens) that support the findings of this study are available in the Scale insect repository of the entomological museum Universidad Nacional Agronomia Bogota – UNAB, Facultad de Ciencias Agrarias, Colombia. All data generated or analyzed during this study are included in this published article and its supplementary information files.

    Acknowledgments

    The authors thank Dr. Andrea Ramos-Portilla for clarifying some aspects of the morphological variations of Rhizoecus species and Dr. Penny Gullan (Australian National University, Canberra, Australia) for reviewing an earlier version of the manuscript. Many thanks to Erika Valentina Vergara (AGROSAVIA) and Dr. Francisco Serna (Universidad Nacional de Colombia) for their help to access the Museum UNAB. Special thanks to Dr Giuseppina Pellizzari (University of Padova, Italy) for advice on scientific nomenclature. This study was financed by Colciencias (Programa Nacional de Ciencias Básicas [National Program on Basic Sciences]), code 110165843233, contract FP44842-004-2015), by the entomological museum UNAB (Facultad Ciencias Agrarias, Universidad Nacional de Colombia, sede Bogotá) and by Federación Nacional de Cafeteros.

  • The authors declare that they have no conflict of interest.

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    Cetinkaya N, Ic E, Erel Y, Ercin D, Ozvatan S, et al. 2023. Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay. Food Materials Research 3:32 doi: 10.48130/fmr-0023-0032
    Cetinkaya N, Ic E, Erel Y, Ercin D, Ozvatan S, et al. 2023. Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay. Food Materials Research 3:32 doi: 10.48130/fmr-0023-0032
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Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay

  • 1.

    Ondokuz Mayıs University, Faculty of Veterinary Medicine, 55200, Samsun, Türkiye

  • 2.

    Turkish Energy, Nuclear and Mineral Research Agency, Nuclear Energy Research Institute, 06980, Ankara, Türkiye

  • Received: 16 June 2023
  • Accepted: 08 September 2023
  • Published online: 10 November 2023
Food Materials Research  3,  Article number: 32  (2023)  |  Cite this article

Abstract: The objective of the present study was to estimate applied doses of irradiated anchovy (1.83−4.22 kGy) and bluefish (1.98−5.40 kGy) for shelf-life extension by using image analysis in combination with comet assay for trading irradiated fish, especially for public health concerns. Comet assay was carried out considering the EN 13784 protocol. Observed comets were evaluated and recorded by the use of image analysis software (BS 200 ProP). Comets with long tails were observed in the irradiated anchovy and bluefish tissues. However, limited numbers of comets in intact cells were determined in the unirradiated samples. The obtained results clearly indicated that head diameter, head DNA, tail DNA, tail length, and tail moment parameters showed very good correlation (R2 = 0.82 and 0.77 for anchovy and bluefish respectively) with the applied doses of irradiation. Using image analysis in combination with comet assay may be a very useful tool for quantification of absorbed doses for local market checks and also in imported irradiated fish for official control at customs.

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    Introduction

    • Anchovy and bluefish are widely consumed and very popular in Türkiye. Considering total fishery production in Türkiye, its volume was about 630,000 tones and shared as: marine capture, 51%; aquaculture, 44% and inland capture, 5% in 2017[1]. The most important habitats of anchovies (commonly, 12−15 cm in length) are the Eastern North and Central Atlantic, Black and Azov Seas. Bluefish (approximately, 20−60 cm in length) are a vigorous, fast, greedy predator, attacking other fish such as anchovies and this fish species finds a habitat in tropical and subtropical seas such as the Western and Eastern Atlantic Ocean, the Mediterranean and the Black Sea[2]. Sea food is extremely perishable products with a narrow shelf life. However, several days lengthening of the shelf life leads to the industry's viability and marketing capacity of sea food[3]. It is recommended that these products can be irradiated at doses of up to 3 kGy which results in a marked drop (app. 2−5 Log10 reduction) of vegetative bacterial pathogen count in fish[4]. During this process, the type of radiation and the energy level, as well as the composition, physical condition, temperature and atmospheric conditions of the food are seen as the main factors determining the effectiveness of the application. While primary radiolysis effects cause chemical changes, highly reactive intermediates undergo various reactions to form stable chemical products. Chemical changes in living materials, on the other hand, produce biological results[5]. Following ionization, physical injury and/or chemical changes occur first, followed by DNA damage, which results in cell death (early effects) and sublethal cellular changes (genetic effects and cancer), respectively[6]. According to Zanardi et al.[7] the flavor-related secondary modifications in irradiated food are those that occur. Lipid oxidation, the emergence of mercaptans, and the depletion of antioxidant vitamins C and E are all factors that contribute to this outcome. The primary radiolytic byproducts include certain cholesterol oxides and furans, as well as 2-alkylcyclobutanones (2-ACBs) made from the principal fatty acids in foods.

      Comet assay or single cell gel electrophoresis assay (SCGE) has become an adaptable method to detect DNA damage in cells and tissues since its ability to measure DNA damage in the form of strand breaks generated by the action of different processes such as irradiation, due to its various advantages, including sensitivity, speed, simplicity, and cost effectiveness[8−11]. Specifically, the comet assay name is derived from the detection of DNA strand breaks and the resulting 'comet' formation in cell DNA as a result of various factors and is based on the principle that damaged DNA migrates at a different rate than undamaged DNA during electrophoresis. That is, when a single cell suspension containing damaged DNA embedded in low-melting agarose is subjected to electrophoresis, the damaged DNA resembling the structure of a comet migrates at a different rate away from the nucleoid body containing undamaged DNA[12]. While this situation is seen in irradiated samples, if the untreated samples do not undergo any DNA fragmentation processes, they always contain intact cells that are not identified as 'comets'. Whereas, unharmed cells can not be observed in an irradiated sample, and the shape of the 'comet' can be used as an indicator of the increase in the applied dose value[13]. After intensive research programs, the European Committee of Standardization (CEN) adopted the DNA comet assay as EN 13784 for the detection of irradiated foods -screening method[14]. In recent years, comet assay has been recognized by different authors[15−19] as a valuable method for detection of irradiation in sea foods. Cerda et al.[20] firmly stated that the development of simple and rapid tests to detect irradiated foods will facilitate food control, and, as a result, consumers will be more confident that radiation processing is properly supervised.

      In this context, the DNA comet assay has an important feature as a simple, low-cost, and fast screening test for qualitative detection of various irradiated foods. Therefore, the purpose of this research was to quantify the applied radiation doses by using the DNA comet assay together with image analysis in irradiated anchovies and bluefishes where quantification of DNA comet analysis parameters was not encountered in these fish in previous articles.

    Materials and methods

      Materials

    • Anchovy (Engraulis encrasicolus) samples were obtained in November and bluefish (smaller) (Pomatomus saltarix) in September from the Black Sea coast of Türkiye and transferred to the Nuclear Energy Research Institute (Ankara) on dry ice in a non-air-tight insulated container (cooler). After about 8 h of transport, collected dead fish samples were prepared for irradiation treatments according to the following sampling plan: i) number of treatment groups: five (control and four different irradiation doses); ii) the number of samples for each group: three (bluefish), 30 (anchovy); iii) the number of parallels: three in every treated sample (Table 1). Anchovy and bluefish samples for each dose level were packed in labelled polyethylene bags, and identified with their respective irradiation doses.

      Table 1.  Sampling and treatment plan.

      SampleTreatment
      Number of treatment groupsAnchovyControl, 1.83, 2.85, 3.42, 4.22 kGy
      BluefishControl, 1.98, 3.93, 4.57, 5.40 kGy
      Number of samples for each groupAnchovy30
      Bluefish3
      Number of parallelsAnchovyControl
      1.83 kGy
      2.85 kGy
      3.42 kGy
      4.22 kGy      		3
      3
      3
      3
      3
      BluefishControl
      1.98 kGy
      3.93 kGy
      4.57 kGy
      5.40 kGy      		3
      3
      3
      3
      3

      Sample irradiation

    • Whole anchovy and bluefish samples were irradiated in a gamma cell (60Co, dose rates were 1.29 and 1.32 kGy/h, respectively) at Nuclear Energy Research Institute, Ankara, Türkiye. Harwell Amber 3042 dosimeters were used for the measurement of radiation dose. Absorbed doses were determined in anchovy and bluefish samples with the dose of 1.83, 2.85, 3.42, 4.22 kGy and 1.98, 3.93, 4.57, 5.40 kGy, respectively (Table 1). Just after irradiation, samples were analyzed by DNA Comet Assay[21].

      DNA comet assay

    • Randomly selected 1 g fillet meat slices was transferred into a beaker; 5 mL of ice-cold phosphate-buffered saline (PBS) was added and stirred (5 min, 500 rpm). The formed suspension was filtered through 500 and 200 µm cloth sieves, respectively, and left on ice for about 5 min. The obtained supernatant was used as a cell suspension (100 µL) which was mixed with 1 mL of low-melting agarose (0.8% in PBS). One hundred µL of this mixture was spread on precoated slides. Then, immersion of coated slides was conducted in a lysis buffer (0.045 M TBE, pH 8.4, containing 2.5% SDS) for 2−9 min. Next, electrophoresis was carried out at 2 V/cm for 2 min using a buffer with the same properties but without SDS. Propidium iodide was used to visualize DNA as described in TS EN 13784 TR[21].

      Evaluation of comets by image analysis

    • Observed comets from the DNA comet assay were evaluated by image analysis in terms of the comet parameters to interpret and estimate the applied doses. Prepared slides were examined with a standard transmission microscope (Olympus BX 51) at 20 X and featured/quantified by a digital color video camera (Pixera) with software image analysis (BS200 ProP, BAB Imaging System, Ankara, Türkiye). Applied image analysis software provides automatic and semi-automatic analysis opportunities. Measured comet parameters were available immediately after 3 s[22−24].

    Results and discussion

    • It is well known that ionizing radiation causes DNA breakage[25]. The pattern formed by the DNA and its apparent fragments (comet) depends on the applied radiation dose[15]. The comet assay technique is based on determination of DNA breaks and it has been applied to several meat tissue samples such as fish[15−19]. Previous confirmed that comet assay is a valuable method for the determination of irradiation in seafood. Therefore, the main goals of the current study were to quantify and evaluate some critical parameters of comet assay analysis in the irradiated anchovy and bluefish samples for official market checks for consumer health concerns.

      This research covered estimation of the effect of gamma irradiation in different doses on DNA of anchovy and bluefish by using comet assay and measurements of the values of different comet as quantified with BS 200 ProP, BAB Imaging System software. The generated data is shown in Table 2.

      Table 2.  The mean values of measured parameters of comets derived from irradiated anchovy and bluefish.

      Absorbed dose (kGy)Measured parameters (Mean ± Std. error)
      HDaHDNAbTDNAcTLdTMe
      AnchovyControl47.09 ± 2.0715.92 ± 1.8384.08 ± 1.8342.23 ± 3.6437.05 ± 3.40
      1.8316.35 ± 0.610.70 ± 0.0899.30 ± 0.08192.52 ± 1.72191.68 ± 1.74
      2.8519.00 ± 0.320.80 ± 0.0499.20 ± 0.04212.58 ± 1.34211.31 ± 1.35
      3.4213.36 ± 0.180.26 ± 0.0799.74 ± 0.07209.43 ± 3.13208.94 ± 3.14
      4.2210.77 ± 0.100.13 ± 0.0399.87 ± 0.03202.16 ± 1.34201.88 ± 1.35
      BluefishControl55.85 ± 1.5910.52 ± 0.5489.48 ± 2.0750.55 ± 0.5445.76 ± 2.22
      1.9817.29 ± 0.750.28 ± 0.0499.72 ± 5.39227.88 ± 0.04227.23 ± 5.39
      3.9312.91 ± 0.120.19 ± 0.0299.81 ± 2.15285.21 ± 0.02284.69 ± 2.15
      4.5712.12 ± 2.660.25 ± 0.1899.75 ± 3.15251.32 ± 0.18250.69 ± 3.49
      5.4011.07 ± 0.240.16 ± 0.0299.84 ± 2.13218.35 ± 0.02217.98 ± 2.13
      a HD: Head diameter. b HDNA: Head DNA. c TDNA: Tail DNA% = percent of DNA in the comet tail. d TL: Tail length = length of the comet tail measured from right border of head area to end of tail (micron = pixels). e TM: Tail moment = Tail DNA% × Tail length (DNA% in the tail).

      During analysis, it was found that the cells' thread breaks appeared like the tails of a comet. In the classification made by eye under the microscope before image analysis and based on a morphological basis, the criteria reported by Marín-Huachaca et al.[17] were taken into consideration. After microgel electrophoresis of cells from control and irradiated fishes, observed images of DNA comets were shown in Fig. 1. The measured parameters were obtained as a result of the evaluation of the detected comets with image analysis and are shown in Table 2, Figs 2 & 3.

      Figure 1. 

      Typical DNA comets are from anchovy and bluefish. Electrophoresis conditions: 2 V/cm for 2 min, propidium iodide staining. All slides were examined at a magnification of 20×.

      Figure 2. 

      Changes in measured DNA comet parameters in anchovy samples.

      Figure 3. 

      Changes in measured DNA comet parameters in bluefish samples.

      The visual examinations allowed rapid sorting and characterization since untreated tissue samples generally have intact structure, but irradiated samples demonstrated increased DNA degradation in terms of increased irradiation doses.

      Such as the comet head parameter, the important parameters in fish samples were markedly changed depending on the dose. Measurement of fragmented DNA migration from the nucleoid body determines its comet head parameters[26]. Increased irradiation doses for anchovy samples resulted in a decrease in head diameter and head DNA parameters from 47.09 to 10.77 µm and from 15.92 to 0.13 µm, respectively. These parameters in bluefish samples declined from 55.85 to 11.07 µm and from 10.52 to 0.16 µm, respectively. Furthermore, tail length and tail moment are commonly used parameters[27]. The tail length (TL) expresses the difference between the greatest movement size of a single tail and the diameter of the comet's head. Furthermore, this parameter reveals the change in tail morphology very clearly. The rotational fluorescence intensity in the tail can be expressed accurately as the tail moment (TM)[28].

      The percentage of DNA in the comet's tail (Tail DNA) is another commonly used parameter for DNA damage. According to Collins et al.[29], 1 Gy of X- or γ-irradiation induces 0.31 breaks per 109 Daltons of DNA. Similar results were found when comparing doses of the detected DNA with the comet assay. Considering our results are related to these parameters in irradiated fish tissues, a significant dose-dependent increase was noticed in tail DNA%, tail length and tail moment parameters (Table 2, Figs 2 & 3).

      The head diameter and tail moment parameter measurements were gathered after exposure to different levels of radiation, correlated with the radiation dose and the square of the correlation coefficient (R2) values calculated for the parameters. Dose dependent response curves of samples were shown in Figs 4 & 5.

      Figure 4. 

      Dose dependent response curve of head diameter and tail moment for anchovy.

      Figure 5. 

      Dose dependent response curve of head diameter and tail moment for bluefish.

      The ratio of the density of the tail of the comet to the head of the comet reveals DNA damage[27]. The tail moment was significantly increased in anchovy and bluefish samples in a dose-dependent manner in response to radiation, with R2 values of 0.74 and 0.65, respectively. Furthermore, the R2 values of the determined Head Diameter parameters were 0.83 and 0.77, respectively. They showed a linear dose-response relationship with DNA damage in increased dose ranges.

      Quantification of comets was carried out by image analysis to evaluate changed parameters in terms of exposed doses. First of all, the results revealed that no comets in untreated fish samples were determined since there was no DNA damage at 0 kGy as shown in Fig. 1. Determined comets were compared to irradiated ones with a round shape, a large and intense head and a short tail in untreated tissues. Whereas, the results of irradiated anchovy (1.83, 2.85, 3.42, 4.22 kGy) and bluefish (1.98, 3.93, 4.57, 5.40 kGy) samples all showed comets depending on DNA damage (Fig. 1). In a previous study, irradiated (0, 2.5 and 5 kGy) and cold stored (2 °C) salmon samples were analyzed by Cerda[15]. One day after irradiation, the typical pattern of irradiated cells (2−3 times longer than comets compared to untreated ones) was determined. Secondly, a decrease in head diameter and head DNA parameters suggests that DNA damage occurred after an increased irradiation dose. Some DNA comet assay parameters investigated in a wide range of radiation doses (0.25, 0.50, 1, 3, 5, 7, and 9 kGy) for rainbow trout samples and reported that proportionally with increasing irradiation dose, the comet's tail length increased while the amount in the comet's head decreased[18]. Thirdly, tail moment also changed exponentially with increasing doses in both anchovy and bluefish R2 = 0.74, R2 = 0.65, respectively. Head diameter values of samples declined linearly R2 = 0.83, R2 = 0.77, respectively (Figs 2 & 3). These results revealed that irradiation treatments up to 5 kGy induced a significant increase in DNA damage in treated fish samples. In another study, the fish samples (salmon, sardine, halibut, herring, plaice, saithe and squid) were irradiated (0.5−5 kGy and in the presence of ice) and stored (−20 °C and 1, 3 and 7 d).

      Khan et al.[16] stated that this assay could be used properly for determining radiation processing at low doses, such as 0.5 kGy for salmon fish. On the other hand, this analysis has failed for halibut, herring, plaice (due to fast natural degradation of DNA), saithe and squid (since there is nonisolation of DNA material).

      Consequently, radiation-processing leads to an extensive molecular size reduction of DNA in treated samples. Our findings are consistent with previous DNA comet studies on animal-origin foods, specifically different fish samples.

    Conclusions

    • The identification of irradiated foods using the analytical method is considered as an important tool in terms of both national and international market control. It permits consumers to buy irradiated foods for eating that have been inspected with oversight by public health agencies. This paper covered the DNA comet assay with image analysis that was carried out to reveal DNA damage levels in anchovy and bluefish samples at applied irradiation doses. Regarding the identification of critical assay parameters (head diameter, head DNA, tail DNA%, tail length and tail moment) and the interpretation of results, irradiation treatment up to 5 kGy induced a significant increase in DNA damage in fish meat. Radiation treated fish tissues were easily distinguished from unirradiated (control) samples after irradiation. Therefore, it is highly feasible that irradiated fish samples will be determined by this screening technique together with image analysis and it can be put into application as a precise routine checking method. As a result, it is fundamentally preferable to rely on a solid and proven, already configured analysis system.

    Author contributions

    • The authors confirm contribution to the paper as follows: study conception and design: Cetinkaya N, Erel Y, Ercin D, Ozvatan S, Yazici N; data collection: Erel Y, Ercin D, Ozvatan S, Yazici N, Ic E; analysis and interpretation of results: Yazici N, Ic E; draft manuscript preparation: Cetinkaya N, Ic E. All authors reviewed the results and approved the final version of the manuscript.

    Data availability

    • To be supplied by author.

      Acknowledgments

      • The authors gratefully thank the Turkish Energy, Nuclear and Mineral Research Agency, Nuclear Energy Research Institute for the financial support (Project No. A3.H1.P5.01).

      Conflict of interest

      • The authors declare that they have no conflict of interest.

      Rights and permissions
      • Copyright: © 2023 by the author(s). Published by Maximum Academic Press on behalf of Nanjing Agricultural University. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (5)  Table (2) References (29)
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    Cetinkaya N, Ic E, Erel Y, Ercin D, Ozvatan S, et al. 2023. Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay. Food Materials Research 3:32 doi: 10.48130/fmr-0023-0032
    Cetinkaya N, Ic E, Erel Y, Ercin D, Ozvatan S, et al. 2023. Estimation of the applied doses in irradiated anchovy and bluefish for shelf-life extension using image analysis in combination with DNA comet assay. Food Materials Research 3:32 doi: 10.48130/fmr-0023-0032
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