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Figure 2 in Spectral characterization and biological evaluation of biomolecules from the peels of three orange fruits: a comparative study
Figure 2. The morphology of orange fruits. (A) Valenica orange; (B) Madarin orange; (C) African Navel orange.
Figure 5 in Spectral characterization and biological evaluation of biomolecules from the peels of three orange fruits: a comparative study
Figure 5. The structure of bioactive compounds of RM of Mandarin orange peel (1) Lim-onene; (2) Octadecanoic acid, 2-hydroxy-1- (hydroxymethyl) ethyl; (3) Hexadecanoic acid, 2-hydroxy-1-(hydroxymethyl) ethyl; (4) Tetradecanamide; (5) n-Hexadecanoic acid; (6) α-D-Mannofuranoside, 1-O-(10-undecenyl)-; (7) 3-Deoxy-d-mannoic lactone; (8) Desulpho-sinigrin; (9) 2-Methoxy-4-vinylphenol; (10) Decanal; (11) Vitamin E; (12) 1-Monolinoleoylglycerol trimethylsilyl ether.
Figure 4 in Spectral characterization and biological evaluation of biomolecules from the peels of three orange fruits: a comparative study
Figure 4. The structure of bioactive compounds of RM of Valencia orange peel (1) Limonene; (2) 9-Octadecenamide, (Z)-; (3) Hexadecanoic acid, 2- hy-droxy-1-(hydroxymethyl)ethyl; (4) Octadecanoic acid, 2-hydroxy-1-(hydroxymethyl)ethyl; (5) Tetradecanamide; (6) Hexadecanamide; (7) Ethyl iso-allocholate; (8) Ethyl α-d-glucopyranoside; (9) d-Glycero-d-galacto-heptose; (10) α-Sitosterol; (11) Vitamin E; (12) 4H-1-Benzopyran-4-one, 2-(3,4-dimethoxyphenyl)-5,6,7-trimethoxy-.
Figure 8 in First information on biology of the needlefish Tylosurus acus imperialis (Belonidae) off the Tunisian coast (Central Mediterranean)
Figure 8. - Growth curve and back calculated length at age by sex of T. a. imperialis from Tunisian coast.
Figure 2 in First information on biology of the needlefish Tylosurus acus imperialis (Belonidae) off the Tunisian coast (Central Mediterranean)
Figure 2. - Length-weight relationships of T.a. imperialis from Tunisian coast. A: Females; B: Males.
Figure 3 in Reproductive biology of Lophius budegassa (Lophiidae) in the North Aegean Sea
Figure 3. - Monthly percentage of each maturity stage for males (A) and females (B) black-bellied anglerfish. Maturity stages: I, Immature; II, Maturing; III, Mature; IV, Spawning; V, Post-Spawning.
Figure 2 in Reproductive biology of Lophius budegassa (Lophiidae) in the North Aegean Sea
Figure 2. - Length-number distribution of Lophius budegassa sampled in the Saros Bay by commercial vessel between September 2006 and September 2008.
Figure 4 in Reproductive biology of Lophius budegassa (Lophiidae) in the North Aegean Sea
Figure 4. - Monthly variation of gonadosomatic index for males (A) Figure 5. - Percentage of sexually mature males (A) and females and females (B) Lophius budegassa between September 2006 and (B) Lophius budegassa according to total length. September 2008. Vertical bars represent standard error with sample size values indicated between brackets. and GSI values in females and males. Therefore, the results suggest a main spawning period from December to March decreases with increasing depth that may reflect a seasonal for blackbellied anglerfish. In females, spawning and postsegregation (García-Rodríguez et al., 2005). Laurenson et spawning stages have been seen from December to March al. (2008) found that the proportion of L. piscatorius varies and in males from January to March (Fig. 3A, B). At the same with both depth and season in Scottish waters. Several state- time, GSI values in males were high except in August and ments can explain this difference. Differentiation in growth November, while female GSI values were the highest in Janrate between sexes can cause an unbalanced proportion, uary (Fig. 4A, B). The spawning males can be seen almost all since the sex presenting faster growth rate will go through year around (Duarte et al., 2001; Landa et al., 2014), and this the most vulnerable smaller size phase quickly and, there- could increase the chances for a mature female to encounter fore decrease the predation proportion. Conversely, the sex with slower growth rate will be more likely to pass predation, with its abundance decreased disproportion- Table I. - Length at 50% maturity (L50) of Lophius budegassa obtained by other authors. ately in next development phases (Vicentini and Araújo,
Fig. 1 in Effect of Different Thermal Conditions on Biology and Number of Generations of Palpita forficifera (Lepidoptera: Crambidae)
Fig. 1. Survival curves for female (A) and male (B) survival of Palpita forficifera at different temperatures (10, 15, 20, 25, and 30 °C), 60 ± 10% relative air humidity, and 14:10 h (L:D) photoperiod. Curves followed by the same letters for each gender did not differ from one another by the log-rank test (Tms = mean time of survival).
Fig. 3 in Effect of forest microhabitat and larval stage on overwintering survival, development, and phenology of Spathius galinae (Hymenoptera: Braconidae), biological control agent of emerald ash borer, Agrilus planipennis (Coleoptera: Buprestidae)
Fig. 3. Proportion of dead (A) and diapaused (B) Spathius galinae by stage at time of deployment, and overwintering microhabitat. Fate was determined by dis- secting all logs once emergence was complete. Letters of the same type and case within the same subfigure indicate significance when data are considered by stage alone (P <0.05).
Fig. 2 in Effect of forest microhabitat and larval stage on overwintering survival, development, and phenology of Spathius galinae (Hymenoptera: Braconidae), biological control agent of emerald ash borer, Agrilus planipennis (Coleoptera: Buprestidae)
Fig. 2. Deployment jar for logs containing emerald ash borer larvae parasitized by Spathius galinae. Logs were inserted in floral foam in 3.8 L polyethylene terephthalate jar with 2 mesh cutouts for ventilation and excess water drain- age. The jar was attached to the tree by resting the bottom of the jar on 2 nails hammered into the tree while a length of wire wrapped around the 2 nails on either side of the jar. Another wire looped around the neck of the jar and was fastened to the nail at the top. Water was added to the jars as needed to ensure adequate hydration of the logs and larvae.
Fig. 2 in Effect of tenthredinid leaf miner invasions on growth of Alaska white birch in Anchorage, Alaska, USA, and the interaction with biological control of amber-marked birch leaf miner
Fig. 2. Time series analysis of mean radial growth increments (mm) of Alaska white birch (Betula neoalaskana) in Anchorage, Alaska (n = 99) from 1984 to 2018: (1) pre-amber-marked birch leaf miner (Profenusa thomsoni) outbreak control period (1984–1995) (black dots); (2) amber-marked birch leaf miner outbreak period (1996–2007) (red dots); and (3) the amber-marked birch leaf miner suppression period due to biological control (2008–2018) (green dots). Time Series Mean = 1.7938, Std = 0.3843, N = 35, Zero Mean ADF (Augmented Dickey Fuller test) = −0.9887, Single ADF = −2.8315, Trend ADF = −4.8800.
Fig. 1 in Effect of tenthredinid leaf miner invasions on growth of Alaska white birch in Anchorage, Alaska, USA, and the interaction with biological control of amber-marked birch leaf miner
Fig. 1. Percentage of Alaska white birch (Betula neoalaskana) leaves in Anchorage mined by the amber-marked birch leaf miner (AMBLM on graph) (Profenusa thomsoni) or the late birch leaf edge miner (LEM on graph) (Heterarthrus nemoratus) from 1990 to 2019, from the initial invasion of amber-marked birch leaf miner (around 1991) through its suppression by classical biocontrol (2004– 2015) and the invasion of a second species of leaf miner (H. nemoratus) (around 2008). Data on percentage of birch leaves mined by each species were taken from multiple sources: (1) P. thomsoni: 2006–2011 (Soper et al. 2015); 2015 and 2018 (Wenninger unpublished 2018); and 2019 (Andersen et al. 2021); and (2) H. nemoratus: 2008–2010 (Lundquist et al. 2012); 2011 (Mulvey &Lamb 2012, p. 15); 2015 and 2018 (Wenninger unpublished 2018); and 2019 (Andersen et al. 2021).
Fig. 1. Experimental microhabitats near the USDA-ARS Louis A in Effect of forest microhabitat and larval stage on overwintering survival, development, and phenology of Spathius galinae (Hymenoptera: Braconidae), biological control agent of emerald ash borer, Agrilus planipennis (Coleoptera: Buprestidae)
Fig. 1. Experimental microhabitats near the USDA-ARS Louis A. Stearns Laboratory in Newark, Delaware, USA. Letters indicate habitat type and approximate experiment locations: (A) mature forest, a larger, more mature wooded area; (B) urban forest, small, highly disturbed woodlot.
Fig. 1 in Does Megaselia scalaris (Diptera: Phoridae) have potential as a biological control agent of fall armyworm?
Fig. 1. Megaselia scalaris (Diptera: Phoridae): adult female (A); eggs (B); larvae dorsal view (C); larvae ventral view (D); and pupae (E).
Fig. 3 in Effect of tenthredinid leaf miner invasions on growth of Alaska white birch in Anchorage, Alaska, USA, and the interaction with biological control of amber-marked birch leaf miner
Fig. 3. Trends in annual radial growth increment (mm) from Alaska white birch (Betula neoalaskana) in Anchorage, Alaska (n = 99 cores) during each of 3 periods: (A) the pre-amber-marked birch leaf miner (Profenusa thomsoni) outbreak period (1984–1995, (B) the amber-marked birch leaf miner outbreak period (1996– 2007), and (C) the biological control amber-marked birch leaf miner suppression period (2008–2018).
Fig. 4 in Effect of forest microhabitat and larval stage on overwintering survival, development, and phenology of Spathius galinae (Hymenoptera: Braconidae), biological control agent of emerald ash borer, Agrilus planipennis (Coleoptera: Buprestidae)
Fig. 4. Survival analysis of Spathius galinae emergence from urban (A) and mature forest (B) sites over time by stage at time of deployment.
Fig. 2 in Erratum: Biological studies of the Oligonychus litchii (Trombidiformes: Tetranychidae) on four commercial litchi cultivars (Florida Entomologist (2019) 102:2 (418-424) DOI: 10.1653/024.102.0220)
Fig. 2. Age-stage-specific survival rate of O. litchii reared on different litchi cultivars at 25 ± 1 °C, 65 to 80% RH, and a photoperiod of 14:10 h (L:D).
Fig. 1 in Erratum: Biological studies of the Oligonychus litchii (Trombidiformes: Tetranychidae) on four commercial litchi cultivars (Florida Entomologist (2019) 102:2 (418-424) DOI: 10.1653/024.102.0220)
Fig. 1. Life stages of Oligonychus litchi: (A) egg; (B) larva; (C) protonymph; (D) deutonymph; (E) adult male; (F) adult female.
Fig. 3 in Erratum: Biological studies of the Oligonychus litchii (Trombidiformes: Tetranychidae) on four commercial litchi cultivars (Florida Entomologist (2019) 102:2 (418-424) DOI: 10.1653/024.102.0220)
Fig. 3. Age-specific survival rate (lx), age-specific fecundity (mx) of O. litchii reared on different litchi cultivars at 25 ± 1 °C, 65 to 80% RH, and a photoperiod of 14:10 h (L:D).
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These curated guides explain access requirements, typical timelines, costs, and reuse considerations for widely used research datasets.
Allen Brain Atlas
Allen Brain Atlas is an Allen Institute collection of brain map atlases, datasets, APIs, and analysis tools covering mouse, human, and non-human primate brain resources.
Annotated Behaviour and Observability Dataset (ABODe)
ABODe is a University of Edinburgh DataShare dataset for behavior classification in group-housed mice using home-cage video, identities, bounding boxes, ground-plate positions, and annotator labels.
DANDI Archive for NWB datasets
DANDI is a BRAIN Initiative archive for publishing and sharing neurophysiology data, including electrophysiology, optophysiology, and behavioral data packaged as NWB and related standards.
International Brain Laboratory public data
The International Brain Laboratory public data releases expose standardized mouse decision-making experiments, including Neuropixels recordings, widefield calcium imaging, behavior, and session metadata accessed through the ONE API.
OpenNeuro
OpenNeuro is a free, open platform for sharing neuroimaging datasets, with public search, dataset pages, and download paths for web, S3, DataLad, and the OpenNeuro CLI.