Salinity is one of the major environmental constraints that has negative effects on plant growth and development. Globally, approximately 800 million hectares of the Earth's land are salt-affected, accounting for around 6% of the total land area, and this is expected to increase in the near future (Abdel Latef et al., 2020;Ruiz-Lozano et al., 2012). The consequences of soil salinity as a major environmental problem are evident in arid and semi-arid regions worldwide (Ahmad et al., 2015;Liu et al., 2016;Osman et al., 2021). Plants grown in salty environments tend to accumulate high levels of salt, which cause disturbances within the physiological and biochemical parameters, leading to ion toxicity, reduced photosynthesis, free radicals' accumulation, instability of membranes, and a variety of metabolic disorders (Atzori et al., 2017;Hanin et al., 2016;Osman et al., 2021). The damage triggered by salinity is influenced by many factors, of which salt concentration, crop sensitivity, and management control tools are the most important (Carillo et al., 2019;IUCN Red List categories and criteria, version 3.1, second edition, 2012;Rouphael et al., 2012). The adaptive strategy of salt-tolerant plants in response to salinity stress takes place by different mechanisms, including osmoregulation, antioxidant defense system, and toxic ion (Abdel Latef et al., 2020). Therefore, studying plant mechanisms under salinity stress is essential to determine whether plants can perform physiological and biochemical processes to cope with salinity stress. Furthermore, the use of sustainable agricultural tools to enhance plant growth and productivity under salt stress can ensure the establishment of high-quality crops under anticipated climate change. It is important to identify environmentally safe and sustainable approaches to mitigate the harmful effects of salinity on plants. One of the strategies to mitigate salt stress is the use of natural plant extracts instead of chemical solutions, for limiting soil, water, and environmental pollution. Natural plant extracts are effective in regulating metabolism, thereby promoting plant growth and yield.Halophytes distributed along the Red Sea and the Mediterranean Sea coastal lands could be used as a valuable source of bio-stimulants for mitigating salinity stress (Osman et al., 2021). Among them, Halocnemum strobilaceum (Pall.) M. Bieb is of particular interest due to its remarkable salt tolerance and adaptive mechanisms that allow it to survive and reproduce in highly saline environments. This small succulent shrub belonging to the family Amaranthaceae is capable of germinating at salinity levels up to 0.5 M NaCl, (Qu et al., 2008) reflecting its efficient osmoprotective and ion-regulation systems. Furthermore, H. strobilaceum is known to possess strong antioxidant potential, with its ethyl acetate extract and flavonoid components exhibiting high radical-scavenging activity comparable to that of standard antioxidants such as Trolox (Radwan and Shams, 2007). The plant's volatile oil is also rich in hydrocarbons and oxygenated compounds, suggesting potential bioactivity. Owing to these characteristics, H. strobilaceum represents a promising candidate for developing eco-friendly biostimulants aimed at enhancing the resilience of salt-sensitive crops like quinoa.Quinoa (Chenopodium quinoa Willd.) is one of the most important economic plants, belonging to the Amaranthaceae family. Its native range is Ecuador to northwestern Argentina, and it is introduced to many countries across the globe. Quinoa grains are highly healthful due to their high protein content and numerous vitamins and minerals. It is also used as a medicinal plant, treating some medical disorders. Quinoa's ability to produce high-protein grains under ecologically extreme conditions makes it important for the variations of future agricultural systems, especially in saline areas of arid lands (Bhargava et al., 2006). Therefore, the main aim of our study is to evaluate the potential cultivation of quinoa plants in saline soils and to assess the effect of H. strobilaceum extract as a bio-stimulant on growth and yield-related traits of quinoa under saline conditions by estimating growth traits, physiological attributes, some enzymatic activities, and protein patterns in salinized quinoa plants. To the best of our knowledge, no study has evaluated the foliar application of H. strobilaceum extract on quinoa under salt stress. The present study explores the potential of a naturally salt-tolerant halophyte as a sustainable source of bioactive compounds that may support quinoa performance under saline conditions, linking native halophyte resources with the improvement of stress resilience in cultivated crops.The halophyte plant material was collected in September 2024 from its natural habitat at Ras El-Hekma shore, Mediterranean Coast, Egypt, 31.225543°N, 27.858790°E. Plants were identified by Amal M. Fakhry, Professor of Biodiversity, Department of Botany and Microbiology, Faculty of Science, Alexandria University. Voucher specimens were deposited in the Herbarium of Alexandria University (ALEX) at the Faculty of Science, deposition numbers ALEX 4123. The samples were airdried and ground to a coarse powder. The powder was then added to distilled water in a weight-tovolume ratio of 1:20 (W/V) and placed in a water bath at 85°C for 20 minutes. The fresh extracts were then filtered through double-layered cheesecloth and allowed to cool to room temperature (Salma et al., 2014). The resulting supernatant was taken as 100% H. strobilaceum water extract and diluted to 50% for usage as foliar spraying (Sofowora, 1982).Seeds of quinoa (Miser 1) were obtained from the Faculty of Agriculture, Alexandria University, Egypt. Homogenous seeds of the quinoa plant were sown in pots in a greenhouse of the Botany and Microbiology Department, Faculty of Science, Alexandria University, Alexandria, Egypt, during the winter season of 2024. The greenhouse conditions included average daytime temperatures of 20-25 °C, nighttime temperatures of 15-18 °C, relative humidity between 55-65%, and a natural photoperiod of approximately 10-11 hours of daylight. To prevent contamination, seeds were sterilized for two minutes in a 1% sodium hypochlorite solution while being shaken constantly. Ten seeds of quinoa were cultivated in plastic pots (30 cm diameter) filled with 7 kg of sandy soil. The experiment was conducted using a two-factor factorial design arranged in a completely randomized design (CRD) with five replicates, where the two factors were salinity levels (0, 25, 50, 75, 100, and 150 mM NaCl) and foliar spray treatments (distilled water and H. strobilaceum extract), resulting in a total of 60 pots. Fourteen days after germination, seedlings were thinned to three per pot, then the pots were irrigated with a constant concentration of salt (0,25,50,75,100,and 150 mM NaCl). The developed plants of both control and NaCl-treated pots were sprayed with H. strobilaceum extract after 14, 21, and 28 days of treatment with NaCl, while the untreated control plants were sprayed with distilled water only. H. strobilaceum extract and NaCl concentrations were selected based on our preliminary experiments. The experimental period extended from 17 November 2024 to 16 February 2025, corresponding to a total duration of 91 days (approximately 13 weeks). Salinity treatment was initiated 14 days after germination and maintained until the end of the experiment through regular irrigation with fixed NaCl concentrations; plant samples were collected after 8 weeks of sowing, while seeds were collected at the end of the experiment after 13 weeks. Plant samples were collected after 8 weeks of sowing to analyze the following criteriaThe shoot height (cm), number of leaves, leaf area (cm 2 ), root length, and dry weight of shoot and root, and seed yield were used as growth parameters. Randomly selected plants from each treatment were used to estimate plant growth parameters at the end of the experiment. The shoot height and root length of quinoa plants were manually measured using a measuring scale. Dry weights for shoot and root were measured after drying samples of fresh weights in the oven at 60 °C until constant weight. The image-processing program ImageJ ver. 1.53t was used to calculate the leaf area in cm 2 (Martin et al., 2020;Schroeder et al., 2021). Additionally, seed yield weight was also assessed after complete growth and plant drying.Chlorophyll fluorescence was measured in fresh leaves using an OS-30P pulse-modulated chlorophyll fluorimeter (Opti-Sciences, Hudson, USA), following the method outlined by Van Kooten and Snel (1990). The maximum quantum efficiency ratio of PSII (Fv/Fm) was determined using an OS-30P pulse-modulated chlorophyll fluorimeter (Opti-Sciences, Hudson, USA).Soluble sugars from dry, powdered quinoa leaves were extracted using a borate buffer (pH 8.5). Following the method of DuBois et al. (1956). Sugar concentration was determined calorimetrically by mixing 0.1 ml of the borate extract with 3 ml of concentrated H₂SO₄ and 1 ml of 5% phenol. The mixture was incubated at 60 °C for 20 minutes, then cooled, and the absorbance was measured at 490 nm. A glucose standard curve was used to calculate the total soluble sugar content (mg/g dry matter).Total protein content was determined following the method described by Pomory (2008). The working reagent was prepared by mixing 1% CuSO₄•5H₂O, 2% sodium potassium tartrate, and 2% Na₂CO₃ in a 1:1:100 ratio. A 0.1 mL aliquot of the extract was added to 3 mL of the working solution, briefly vortexed, and left to stand for 10 minutes. Then, 0.1 mL of 1 N Folin-Ciocalteu reagent was added, and the mixture was vortexed again. minutes, the absorbance was measured at nm. content was as of per of dry material content was determined using the method described by and with as the 0.1 of powdered leaf was with 10 mL of and the resulting supernatant was used for A 0.1 mL aliquot of the plant extract standard was with mL of a of 0.5 mL of 1% in 0.5 M buffer (pH mL of solution, and mL of 0.5 M The mixture was shaken and in a water bath at a temperature for minutes, then mixing the absorbance was measured at nm. content was as dry based on a content was determined following the method of et al. A 0.1 of was in mL of and the was to One of the supernatant was with 2 mL of by of in mL of and 20 mL of M 2 mL of The mixture was incubated at °C for 1 and the was then by the in an The resulting was extracted with mL of to room and its absorbance was measured at nm. concentration was as dry using a standard curve prepared with a was following the method of and A 0.1 of leaf was in mL of 5% and at for 10 minutes. The mixture for included 2 mL of 2% sodium 2 mL of N 1 mL of mM and 1 mL of the The mixture was incubated in a water bath at 60 °C for minutes, then and at for 10 minutes. was at nm. content was as dry using a standard curve prepared with content was using the method of and A 0.1 of was extracted three with and the resulting were and to a of 10 One of this extract was with 0.1 mL of reagent and 1 mL of then diluted to mL with distilled minutes, the absorbance was measured at nm. content was as dry using a standard curve prepared using the content in quinoa plants was measured following the method of et al. A 0.1 of leaf was in mL of and at for minutes. 0.5 mL of the supernatant was with 0.5 mL of 10 mM potassium buffer (pH and 1 mL of 1 M potassium The absorbance of the mixture was at nm. content was using an of and as per fresh in quinoa leaves was assessed by measuring the concentration of a of the of using the method of and A 0.5 of fresh leaves was in 10 mL of 5% and the mixture was at for minutes. of the resulting supernatant were with 2 mL of The mixture was incubated in a water bath at °C for 20 minutes, then was measured at and for at nm. concentration was using an of and as per of fresh of dry, powdered leaf samples was using a mixture of and in a ratio. 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Aljeddani et al. (Mon,) studied this question.