Perithecial formation in Gelasinospora reticulispora III. Inhibitory effects of near-UV and blue light during the inductive dark period

Growing hyphae of Gelasinospora reticulispora required a continuousdark period prior to photoinduction of perithecia. The inductivedark period was interrupted by brief exposure of the hyphaeto white light so that the formation of perithecia no longertook place. Photosensitivity of the hyphae in terms of the light-breakeffect gradually changed during the inductive dark period. Sensitivityreached its maximum at the 18th hr of the dark period when anirradiation of 1?10⁵ ergs cm^–2 of near-UV light or 4?10⁴ergs cm^–2 of blue-light was sufficient for the light-break.Red and far-red light had no effect at all. The light-breakeffect was limited to the irradiated portion of the hyphae anddid not affect any unirradiated portions. Inhibitory effecton perithecial formation of continuous white light could betotally replaced for several days with intermittent irradiationof near-UV or blue light if given for 5 min every 4 hr. (Received December 18, 1973; )     

A Versatile Stain for Pollen Fungi, Yeast and Bacteria

A versatile stain has been developed for demonstrating pollen, fungal hyphae and spores, bacteria and yeasts. The mixture is made by compounding in the following order: ethanol, 20 ml; 1% malachite green in 95% ethanol, 2 ml; distilled water, 50 ml; glycerol, 40 ml; acid fuchsin 1% in distilled water, 10 ml; phenol, 5 g and lactic acid, 1-6 ml. A solution has also been formulated to destain overstained pollen mounts. Ideally, aborted pollen grains are stained green and nonaborted ones crimson red. Fungal hyphae and spores take a bluish purple color and host tissues green. Fungi, bacteria and yeasts are stained purple to red. The concentration of lactic acid in the stain mixture plays an important role in the differential staining of pollen. For staining fungi, bacteria and yeasts, the stain has to be acidic, but its concentration is not critical except for bacteria. In the case of pollen, staining can be done in a drop of stain on a slide or in a few drops of stain in a vial. Pollen stained in the vial can be used immediately or stored for later use. Staining is hastened by lightly flaming the slides or by storing at 55±2 C for 24 hr. Bacteria and yeasts are fixed on the slide in the usual manner and then stained. The stock solution is durable, the staining mixture is very stable and the color of the mounted specimens does not fade on prolonged storage. Slides are semipermanent and it is not necessary to ring the coverslip provided 1-2 drops of stain are added if air bubbles appear below the coverslip. The use of differentially stained pollen mounts in image analyzers for automatic counting and recording of aborted and nonaborted pollen is also discussed.     

营养液强度对AM真菌生长发育的影响*

研究了在贫营养基质中不同强度Hoagland营养液对丛枝菌根(Arbuscular mycorrhizae, M)真菌Glomus versiforme生长发育的影响。结果表明:本试验条件下,菌根侵染率、菌丝量、孢子数间呈显著正相关。在施加5%~50%强度Hoagland营养液时, 菌根真菌的生长与宿主植物高粱根中磷浓度、可溶性糖浓度密切相关,而与氮浓度无显著相关。由此认为:在盆栽生产菌根菌剂时,基质中存在一个临界磷浓度,在这个临界浓度之下,菌根真菌的生长发育随磷浓度的提高而增长,超过该临界浓度则会随磷浓度的提高而下降。施用20%、50%强度Hoagland营养液对菌根真菌生长最为有利,其菌根侵染率、菌丝量、孢子数均高于其它处理,因此认为:宿主植物—菌根真菌之间共生关系的基础是营养条件,基质中养分的高低会影响互惠共生关系的建立和发展。在高质量菌剂生产中, 菌根共生体双方的生长发育完全可以由人工控制。施加营养液是一种有效的调控手段,有可能使共生平衡向有利于菌根真菌生长发育的方向倾斜,使真菌得到最大程度的生长。     

Selective staining methods for cartilage of rat fetal specimens previously treated with alizarin red S.

A convenient method for staining cartilage with several basic stains after alizarin red S staining of bone was investigated in rat fetuses. It was found that bromophenol blue was useful and effective for staining of the margin and center areas of cartilage, even in specimens stored in glycerin for over 10 years. The specimens were washed in running tap water for 1 hr, and subsequently were immersed in water or in 70% ethanol at pH 4 for 1 hr or longer. The specimens were then stained with 0.005% bromophenol blue in 40% ethanol adjusted to pH 4 for 2 hr, or with 0.001% bromophenol blue in 40% ethanol adjusted to pH 4 for 24 hr. Furthermore, the bromophenol blue stain color actually faded when the specimens were immersed in water or in 70% ethanol at pH 8. Descending order of the stain-effective action on fetal rat cartilage for the basic stains tested was bromophenol blue, aniline blue, Evans blue, methyl violet, trypan blue, and water blue.     

Brazilin-Toluidine Blue O and Hematoxylin-Darrow Red Methods for Brain and Spinal Cord

Tissue fixed in 10% formalin, formalin-95% ethanol 1:s CaCO₂ or phosphate buffer neutralized formalin, or methanol-chloroform 2:1, was dehydrated and embedded in paraffin or double-embedded by infiltration in 1% celloidin followed by a chloroform-paraffin sequence. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 24-26 C. For either method, mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. For brazilin-toluidne blue O, myelin was stained for 20-60 min, depending upon section thickness, in a self-differentiating solution consisting of: 0.15% Li₂CO₃ 75 ml; 6% brazilin in 95% ethanol, 25 ml; and NaIO₃ 75 mg. After a thorough washing, Nissl material was stained for 3-8 min in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; and 1% toluidine blue 0, 2.5 ml. For hematoxylin-Darrow red, myelin was stained for 2-6 hr in a self-differentiating solution consisting of: 0.15% Li₂,CO₃ 95 ml; 10% hematoxylin in 95% ethanol, 5 ml; and NaIO₃ 25 mg. After a thorough washing, Nissl material was stained for 20 min or less in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; Darrow red, 25 mg. This mixture was first boiled, cooled to room temperature and filtered. In both methods, washing, dehydration, clearing, and mounting completed the process. In the brazilin-toluidine blue technic, myelin sheaths were stained reddish purple; neuronal nuclei light blue with dark granules of chromatin; nucleoli dark blue; and cytoplasm blue with dark blue Nissl granules. In the hematoxylin-Darrow red procedure, myelin sheaths were blue-black; nuclei light red with dark granules of chromatin; nucleoli almost black; and cytoplasm red with bright red Nissl granules.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

Toluidine Blue-Alizarin Red S Staining of Cartilage and Bone in Whole-Mount Skeletons in Vitro

Cartilage and bone of the developing skeleton can be reliably differentiated in whole-mount preparations with toluidine blue-alizarin red S staining after FAA fixation. The recommended staining procedure is based chiefly on the use of newborn white and Swiss-Webster mice, 4-9 days postnatal, but was tested also on mice and rats 3-8 wk of age. Procedure: Sacrifice, skin, eviscerate, remove body fat, and place specimens in FAA (formalin, 1; acetic acid, 1; 70% alcohol, 8) for approximately 40 min. Stain in 0.06% toluidine blue made in 70% ethyl alcohol for 48 hr at room temperature. Use 20 volumes of stain solution to the estimated volume of the specimen. Destain soft tissues in 35% ethyl alcohol, 20 hr; 50%, 28 hr; and 70%, 8 hr. Counterstain in a freshly prepared 1% aqueous solution of KOH to which is added 2-3 drops of 0.1% alizarin red S per 100 ml of solution. Each day for 3 days, transfer the specimen to a fresh 1% KOH-alizarin mixture, or until the bones have reached the desired intensity of red and soft tissues have cleared. Rinse in water, and place in a 1:1 mixture of glycerol and ethyl alcohol for 1-2 hr, then transfer the specimen to fresh glycerol-alcohol for final clearing and storage. Older mice and rats require procedural modifications: (1) fixation for 2 hr, (2) 0.12% toluidine blue, (3) maceration for 4 days in 3% KOH-alizarin, and (4) preliminary clearing for 24 hr in a mixture of glycerol, 2; 70% ethyl alcohol, 2; and benzyl alcohol, 1 (v/v) before placing in a 1:1 alcohol-glycerol mixture.     

Perithecial formation in Gelasinospora reticulispora II. Promotive effects of near-ultraviolet and blue light after dark incubation

Hyphae of Gelasinospora reticulispora growing on corn meal agarat 25?C required light of form perithecia. This response tolight was highly correlated to the length of the preliminarydark period, i.e. the photoinduction of perithecia never occurredin cultures grown in the dark for periods of 27 hr or less,whereas hyphae became photosensitive if incubated for 30 hror longer in the dark. The perithecia became simultaneouslyvisible with the 30 to 48 hr dark-grown hyphae irrespectiveof the different dark periods, but time courses for cultureshaving 54 hr or longer of darkness were dependent upon the timewhen light was given. Light induced perithecia only in the dark-grown portions ofthe hyphae. The longer the dark period, the more the sensitivityto light increased when the most effective range of wavelenghtswas shifted from near-ultraviolet to blue. Light of green andlonger wavelenghts was not at all effective irrespective ofthe duration of darkness. The photoinduced stimulus was notmovable from the irradiated to the unirradiated portions. (Received August 3, 1973; )     

A Modified One-Step Trichrome Stain for Demonstration of Fine Connective Tissue Fibers

Gomori's one-step trichrome procedure was modified to improve coloration of fine connective tissue fibers. Paraffin sections from tissues fixed in alcohol, acetone, Zenkerformol, 10% formalin, Kaiserling's or Carnoy's fluid were mordanted 1 hr at 56 C in Bouin's solution, stained 1 min in a trichrome solution (chromotrope 2R-phosphomolybdic acidaniline blue WS) adjusted to pH 1.3 with HCl, rinsed in 1% aqueous acetic acid, dehydrated and covered. Collagen, reticulum fibers, basement membranes, ring fibers around splenic sinuses, intercalated discs in cardiac muscle and cartilage were colored blue. Nuclei, cytoplasm, fibrin, muscle fibers and elastic fibers were stained red. Pretreatment of sections with Bouin's solution enhanced the affinity of tissues for chromotrope 2R and was found essential for satisfactory coloration of material fixed in alcohol, acetone, formalin or Carnoy's fluid. Because this method does not require differentiation, it gave uniform results even in the hands of inexperienced laboratory trainees. No fading was observed in sections stored for more than 8 yr.     

Wool Fibres: Sectioning and Staining, Differentiation of Ortho and Paracortex

A double embedding technique for tangential sectioning of hair and wool fibres is as follows: The cleaned fibre bundle is attached to a U-shaped, 16 gauge, tinned-copper wire frame with collodion adhesive, soaked in 6% nitrocellulose for 1 hr, and treated with chloroform for 2 hr. The hardened bundle is then cut fom the wire support and embedded in paraffin-beeswax, 95:5. Sectioning is at 6-8 μ. The use of 2% orange G or saturated aqueous picric acid for quantitative study of the fibres, and the demonstration of wool fibre cortical fractions by staining with polychrome methylene blue after oxidation of the sectioned fibres in a solution of formic acid (98/100 w/v) 25 ml; distilled water, 65 ml; and H₂O₂ (30% w/v), 10 ml, for 1 hr, is recommended.     

Sugar production from agricultural woody wastes by saccharification with Trichoderma viride cellulase.

The saccharification of agricultural woody wastes was studied using a commercial enzyme preparation, Cellulase onozuka, derived from Trichoderma viride or the solid culture extracts of the fungus. With the intention of producing sugar at low cost, a simple procedure of enzymatic saccharification of rice straw, bagasse, and sawdust was studied. Delignifying methods of these wastes were investigated using dilute sodium hydroxide solution and dilute peracetic acid. Rice straw and bagasse were effectively delignified by boiling in a 1% sodium hydroxide solution for 3 hr or by autoclaving at 120 degrees C in a 1% sodium hydroxide solution for 1 hr. The sawdust from a broad leaved tree (Machilus thunbergii) was delignified by autoclaving at 120 degrees C in a 1% sodium hydroxide solution for 1 hr and by subsequent boiling in diluted 1/5 peracetic acid for 1 hr. This type of sawdust was also delignified by boiling in 1/5 peracetic acid for 1 hr and by subsequent autoclaving at 120 degrees C in a 1% sodium hydroxide solution for 1 hr. The sawdust from a coniferous tree (Cryptomeria japonica) was delignified by boiling in 1/5 peracetic acid for 1 hr and by subsequent autoclaving at 120 degrees C in a 1% sodium hydroxide solution for 1 hr; however, the successive treatment by autoclaving with alkali solution and subsequent boiling with diluted peracetic acid failed to bring about the desired effect. The saccharification of delignified rice straw, bagasse, and sawdust was examined using Cellulase onozuka, wheat bran or rice straw solid culture at various substrate concentrations, resulting in the formation of 5 to 10% sugar solutions after incubation at pH 5.0, 45 degrees C for 48 hr. The optimum substrate concentration existed at around 10%. Reuse of cellulase solution and resaccharification of residual sawdust were considered to be inadequate.     

The Demonstration of Beta Cells in Pancreatic Islets and Their Tumors

The stain is applied routinely to tissues fixed in 10% buffered formalin (pH near 7.0) or in Bouin's fluid. Bring paraffin section to water as usual and mordant 72 hr in 5% CrCl₃ dissolved in 5% acetic acid. Wash in water and in 70% alcohol and stain 6 hr. Formula of staining solution: new fuchsin, 1% in 70% alcohol, 100 ml; HCl, conc., 2 ml and paraldehyde, 2 ml, mixed together and added to the dye solution; let stand 24 hr before use. After staining, wash in running tap water 5-10 min, rinse in distilled water and counterstain if desired. Dehydration in alcohol, clearing and covering completes the process. When the paraldehyde is obtained from a freshly opened bottle, standardized staining times can be used and thus eliminate the necessity of differentiating individual slides. The granules of beta cells stained deep blue to purple and were demonstrated in the pancreatic islet of man, dog, mouse, frog, guinea pig and rabbit.     

Growth and respiratory activity of mold fungus (Trichoderma lignorum)

The growth ofTrichoderma lignorum was studied in relation to the carbon balance of the culture system and the respiratory activity at different growth phases. Conidia, after inoculation into the medium, swelled and germinated rapidly. The growth rate of the hyphae at the exponential phase was 0.46 hr⁻¹ (2.2 hr for mass doubling) at 25 C. The yield efficiency of hyphal biomass-C at the cost of glucose-C was 67%, while those of the waste products-C excreted and of CO₂-C evolved were, 6.5% and 26.5%, respectively. The yield efficiency of conidia-C to the decrease of hyphae-C was 34%. The germination, growth rate and carbon balance were not affected by different concentrations of glucose from 10 to 2×10³ mg glucose-C/l. Carbon dioxide was needed as the growth factor for the initiation of the germination of conidia, but there was no increase in yield efficiency as a result of CO₂ fixation. The respiratory rate of the fungus changed drastically as the growth proceeded. The rate of endogenous respiration of conidia was less than 0.2 mg CO₂-C/g conidia-C/hr which increased immediately after inoculation into the medium. The highest respiratory rate of hyphae (100–110 mg CO₂-C/g hyphae-C/hr) was obtained throughout the exponential phase. Thereafter, decreasing rapidly, the respiratory rate of submerged hyphae of 1-week-old showed only 1.8 mg CO₂-C/g biomass-C/hr, whereas the rate of aerial hyphae forming conidia increased again, but did not exceed 10 mg CO₂-C/g biomass-C/hr.     

The Demonstration of Beryllium Oxide in Paraffin Sections with Chromoxane Stains

Aqueous solutions of the arylmethane dyes Chromoxane pure blue BLD (C.I. No. 43825) and Chromoxane pure blue B (C.I. No. 43830) will stain beryllium oxide. In the presence of EDTA the staining of other metals is masked. As a specific stain for BeO, formol saline fixed paraffin sections are hydrated and stained for 1 hr with either 0.1 gm of pure blue BLD in 100 ml of pH 4.0 Na-acetate buffer or with 0.1 gm of pure blue B in 1 N NaOH adjusted to pH 9.0 with HCl. To mask interference from other metal ions, 9 gm of Na₂-EDTA is added to 100 ml of the stain solution. BeO is stained blue, organic tissue components are either unstained or pink. Results of tests against other materials show that a high degree of specificity may be expected from these dyes. A 1% aqueous solution of neutral red may be used as a counterstain.     

Luxol Fast Blue Arn: A New Solvent Azo Dye with Improved Staining Qualities for Myelin and Phospholipids

Luxol fast blue ARN (Du Pont, C.I. solvent blue 37) is a diarylguanidine salt of a sulfonated azo dye. This dye was compared with other Luxol blue and Luxol black dyes. Luxol fast blue ARN has improved staining qualities for phospholipids and myelin, and can advantageously be substituted for Luxol fast blue MBS (MBSN). Appropriate staining times for a 0.1% dye solution in 95% ethanol (containing 0.02% acetic add) at 35°-40° C range from 2-3 hr. After staining, the sections should be rinsed in 95% ethanol, rinsed in distilled water, and differentiated for 2 sec in 0.005% Li₂CO₃, rinsed in 70% ethanol, washed in water, and counterstained as required. Phospholipids and myelin selectively stain deep blue. A fixative containing CaCl₂, 1%; cetyltrimethylammonium bromide, 0.5%; and formaldehyde, 10%, in water gave excellent results with brain. However, 10% formalin can be used. The staining of the phospholipids is probably due to the formation of dye-phospholipid complexes.     

Primary fluorescence (Autofluorescence) of fruiting bodies of the wood-rotting fungusFomes fomentarius

Autofluorescence of fruiting bodies of the wood-rotting fungus Fomes fomentarius has been observed and is described among native macrofungi for the first time. The strongest yellow autofluorescence with blue excitation was displayed by pith sets, a weaker yellow, yellow-green to yellow-red fluorescence was due to generative thin-walled hyphae while the weakest yellow-reddish fluorescence was emitted by thick-walled skeletal hyphae (though their parts may emit a more intensive yellow fluorescence). This yellow, yellow-green to yellow-red autofluorescence was assessed to be more intensive than the emission described so far in bacteria and fungi (except for lysed hyphae of the fungus Trametes versicolor). With green excitation all F. fomentarius cells emitted strong red autofluorescence.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Sensory Nerve Endings Shown by Nitro Blue Tetrazolium in Isolated Muscle Spindles of the Cat

Muscle spindles were isolated from freshly removed cat lumbrical muscles in oxygenated Ringer's solution and placed in a solution containing 10 ml of 0.1% nitro blue tetrazolium and 10 ml of 0.2 M phosphate buffered sodium succinate, pH 7.6. Spindles were incubated in this solution at 37 C for 4-12 hr, returned to Ringer's for 30 min at room temperature, fixed in 10% formal-Ringer's for 30 min, and stored indefinitely in distilled water. With this technique the patterns of sensory innervation can be clearly visualized by the deposition of diformazan. The stained preparations may be mounted in glycerol and teased further for whole mount inspection or they may be embedded in Epon and serially sectioned for more detailed study.